Contrasting warming and ozone effects on denitrifiers dominate soil

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Contrasting warming and ozone effects on denitrifiers dominate soil N2O emissions Yunpeng Qiu, Yu Jiang, Lijin Guo, Kent O. Burkey, Richard W. Zobel, H. David Shew, and Shuijin Hu Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.8b01093 • Publication Date (Web): 29 Aug 2018 Downloaded from http://pubs.acs.org on August 30, 2018

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Environmental Science & Technology

Contrasting warming and ozone effects on denitrifiers dominate soil N2O emissions

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Yunpeng Qiu†,//, Yu Jiang†,⊥, Lijin Guo†, $, Kent O. Burkey#, &, Richard W. Zobel&, H. David

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Shew†, and Shuijin Hu†,*

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†

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27695, USA

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//

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210095, China

Department of Entomology & Plant Pathology, North Carolina State University, Raleigh, NC

College of Resources and Environmental Sciences, Nanjing Agricultural University, Nanjing

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⊥

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$

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China

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#

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&

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USA

Institute of Applied Ecology, Nanjing Agricultural University, Nanjing 210095, China College of Plant Science & Technology, Huazhong Agricultural University, Wuhan 430070,

USDA-ARS, Plant Sciences Research Unit, Raleigh, NC 27607, USA Department of Crop and Soil Sciences, North Carolina State University, Raleigh, NC 27695,

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ACS Paragon Plus Environment


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Abstract

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Nitrous oxide (N2O) in the atmosphere is a major greenhouse gas and reacts with volatile organic

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compounds to create ozone (an air pollutant) in the troposphere. Climate change factors such as

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warming and elevated ozone (eO3) affect N2O fluxes, but the direction and magnitude of these

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effects are uncertain and the underlying mechanisms remain unclear. We examined the impact of

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simulated warming (control + 3.6 oC) and eO3 (control + 45 ppb) on soil N2O fluxes in a soybean

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agroecosystem. Results obtained showed that warming significantly increased soil labile C,

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microbial biomass and soil N mineralization, but eO3 reduced these parameters. Warming

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enhanced N2O-producing denitrifers (nirS- and nirK-type), corresponding to increases in both

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the rate and sum of N2O emissions. In contrast, eO3 significantly reduced both N2O-producing

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and N2O-consuming (nosZ-type) denitrifiers but had no impact on N2O emissions. Further, eO3

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offsets the effects of warming on soil labile C, microbial biomass and the population size of

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denitrifiers, but still increased N2O emissions, indicating a direct effect of temperature on N2O

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emissions. Together, these findings suggest that warming may promote N2O production through

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increasing both the abundance and activities of N2O-producing microbes, positively feeding back

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to the ongoing climate change.

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TOC/Abstract art

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INTRODUCTION

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N2O is a potent greenhouse gas with a global warming potential 300 times greater than that of

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carbon dioxide (CO2)1,2 and is the single most important depleter of stratospheric ozone (O3).3 In

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the troposphere, however, N2O reacts with volatile organic compounds to create O3, an air

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pollutant that also contributes to global warming.4 Since industrial times, human activities have

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increased global atmospheric O3 concentration.5,6 Various models predict that global surface

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temperature will be increased by 1-6 oC by the end of this century.1,7 Many regions of the globe

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have experienced surface O3 concentration greater than 40 nmol mol-1 (i.e., 40 ppb), a threshold

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set for injury to O3-sensitive plants.6,8 The current global atmospheric ozone concentration has

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increased from an estimated preindustrial level of 10-15 nmol mol-1 to current ca. 60 nmol mol-

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and chemical N fertilizer applications.1,10 Soil N2O is mainly produced by two microbial

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processes: nitrification (i.e., oxidation of NH4+ into NO3-) and denitrification (i.e., reduction of

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NO3- to gaseous forms, N2O, NO and N2).11-13

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. Globally, agricultural soils account for nearly 70% of the annual N2O budget owing to manure

Climate change factors such as elevated CO2 (eCO2), warming and elevated O3 (eO3) may

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significantly impact N2O emissions.14-16 While eCO2 tended to increase N2O emissions17,18, the

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impact of warming was more variable, with positive,19 negative20 or even neutral effects21 having

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been reported under different conditions.22-24 These inconsistent results may stem from the fact

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that warming impacts multiple processes, some of which may have contrasting effects on soil

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N2O production and consumption.14,21 On one hand, warming may increase N2O emissions

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through directly stimulating the growth and activities of N2O-producing microbes19,25 and/or

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indirectly enhancing the availability of C via plant growth,21,26 and mineral N via mineralization


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(Figure 1).14,27 On the other hand, warming may enhance plant growth, evapotranspiration and N

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uptake, and reduce soil available N21,28 and soil moisture, thereby constraining nitrifiers and

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denitrifiers and their activities.29-31 In contrasting to the direct effect of warming, eO3 affects

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nitrifiers and denitrifiers, and their activities mainly through its indirect effect on plants (Figure

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1).32,33 This is because O3 itself hardly penetrates into soil to directly affect soil microbes.34

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Rather, eO3 often reduces plant growth, plant N acquisition and photosynthate allocation

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belowground for soil microbes,35-37 altering soil N and/or labile C availability for nitrifiers or

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denitrifiers (Figure 1).38-40 However, only a few experiments have so far examined the effect of

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eO3 on N2O emissions.32,39 Negative O3 effect on N2O has been reported in a meadow

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ecosystem41 and in rice fields with the presence42 or the absence of roots,43 suggesting that

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reduced denitrification resulting from O3-induced decreases in plant-derived C inputs may play a

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dominant role.

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An increasing number of experiments have recently examined the impact of warming on

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microbial activities and N2O emission.19,30,44 However, climate warming will likely concur with

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other components such as N deposition and eO3 under future climate change scenarios. Yet, few

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have examined the combined effect of warming and eO3 on N2O emissions. Even fewer have

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attempted to establish the linkages between climate change-induced alteration in N2O emission

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and populations of N2O-producing microbes in field, partially due to the difficulty in quantifying

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N2O–producing microbes.

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Both nitrifiers and denitrifiers contribute to N2O emissions from soil. Ammonia-

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oxidizing bacteria (AOB) and ammonia-oxidizing archaea (AOA) regulate the rate-control step

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of nitrification that produces N2O under aerobic conditions45,46 and their amoA gene that encodes

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the ammonia monooxygenase (AMO) has been effectively used to quantify their abundances in



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soil.47-49 Although denitrifiers are highly diverse heterotrophs and is currently impossible to

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quantify their species composition,50 the function genes encoding nitrite reductase (Nir), the key

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enzyme in the dissimilatory denitrification, have been widely used to characterize the

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abundances and diversity of denitrifiers.50-52 Genes nirK and nirS encode two structurally

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different enzymes catalyzing nitrite reduction: the copper and cytochrome cd1-nitrite reductase,

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respectively.51,53 Another gene, nosZ, which encodes nitrous oxide reductase to convert N2O to

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N2 has also been widely used to describe denitrifier communities.50,54

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We initiated a long-term field experiment in a soybean agroecosystem to understand how

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warming, elevated ozone and their interactions affect plant growth, and soil C and N cycling at

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Raleigh, NC, USA. Taking advantage of this field study, we examined the impact of warming

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and ozone on N2O fluxes, soil labile C and N pools, and the abundances of nitrifiers and

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denitrifiers that are primarily responsible for N2O emissions. Our hypotheses (Figure 1) were that

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(1) warming increases the abundances of N2O-producing microbes and N2O emissions mainly

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through enhancing labile C and mineral N sources rather than through its direct impact on the

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N2O-producing microbes, (2) eO3 reduces the abundances of denitrifiers and N2O emissions by

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reducing organic C inputs, (3) the effects of warming and eO3 on N2O-producing microbes offset

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each other.

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MATERIALS AND METHORDS

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The experimental site

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The long-term experiment was initiated in June 2013 to investigate the response of soybean

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plants to elevated atmospheric O3 and climate warming using an air exclusion system (AES, 3m


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× 10m). The experimental site is located at the Lake Wheeler Experimental Station, 5 km south

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of North Carolina State University, Raleigh, North Carolina, USA (35。43ʹN, 78。40ʹW; elevation

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120 m). The soil is an Appling sandy loam (fine, kaolinitic, thermic Typic Kanhapludult), well

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drained with a pH of 6.2, and contained 12.5 g C and 0.94 g N kg-1 soil when the experiment

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started. Before O3 and warming treatments were initiated, the soil was repeatedly turned-over

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using a disc implement and rotovator.

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Each AES plot consisted of two double-walled open-top chamber panels55 attached to

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aluminum frames and placed in parallel to establish a 3 m x 10 m footprint. A continuous flow of

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air was supplied through the holes in the internal wall of each panel by a fan box containing an

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activated charcoal filter to reduce the O3 concentration of incoming ambient air to ca. 15 nmol

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mol-1. In elevated temperature plots, each fan box was equipped with two 4000W electrical

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resistance heaters (a total of 16000W per plot) and an inline water-to-air heat exchanger. The

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electrical resistance heaters operated 24 hours per day and solar-heated water circulated through

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the heat exchanger during daylight hours. We designed a warming treatment with an increase on

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the ambient air temperature by 3.6 oC. We also selected an elevated ozone concentration at 60

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nmol mol-1, a value close to the average and ozone concentration in our area. We did not increase

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the O3 concentration because the ambient O3 already has harmful effects on some soybean

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cultivars. Moisture was added to the heated air using misters inside the double walled panel to

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elevate the relative humidity to near ambient levels prior to distribution within the experimental

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plot. O3 was generated from O2 by corona discharge (model TG-20, Ozone Solutions, Hull, IA,

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USA). The O3 concentration in each plot was monitored at canopy height with a UV photometric

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O3 analyzer (model 49C, Thermo Environmental Instruments Co., Franklin, MA, USA). Air

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temperature and relative humidity were monitored using HOBO sensors (model U23-001,



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HOBO Pro V2 Temp/RH Datalogger, Onset Computer Corporation, Bourne, MA, USA). Soil

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temperature was measured with an Enviropro dielectric soil probe (model EP100D, Apcos Pty

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Ltd, Adelaide, Australia).

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This experiment was a 2×2 factorial design with four treatments assigned into three

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blocks (replicates). Four different treatments were: (a) charcoal-filtered air and ambient

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temperature (CF); (b) charcoal-filtered air plus 3.6 oC increase in temperature (T); (c) charcoal-

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filtered air plus ambient temperature and 1.4 times ambient ozone (O3);38,39 (d) charcoal-filtered

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air + warming + 1.4 times ambient ozone (T+O3). The purpose of filtration of ambient air with

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activated charcoal was to reduce ambient O3 concentrations to levels considered nonphytotoxic

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to soybeans. The seasonal daily average concentrations (12-h) of O3 and degree of temperature

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over the experimental duration are shown in Table 1.

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Soybean (cultivar Jake) was selected based on our preliminary greenhouse experiment in

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2015 showing that 60 ppb resulted in a moderate foliar ozone response for this cultivar. Soybean

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seeds were planted in the first week of June, 2015 into AES plots. Each plot consisted of eight

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plant rows with a row spacing of 25 cm. During soybean growing seasons, plants were irrigated

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with drip lines to prevent water stress. There were in total three harvesting of aboveground

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biomass in August, September and October and two harvesting of root biomass in August and

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September each year. Aboveground biomass was harvested at soil surface. Three soil cores of 5

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cm in diameter were taken to a 20-cm depth in each plot to harvest three plant roots. Roots were

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washed with tap water. Aboveground and root biomass were weighed after oven drying at 70 °C

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for 72 h.

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Field N2O measurements


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Each AES plot has 8 rows with two static gas chambers randomly installed into all the plots. Gas

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samples were taken within 24-48 hours following an irrigation event approximately every 10

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days. In total, we have 13 gas measurements through the growing season (June to October). The

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chambers (sampling volume is 6392 ml) were closed with a lid, and fitted with a rubber septum

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to allow gas sampling via syringe. Gas samples (5 ml) were taken with 20 ml PE syringes

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(Beceton Dickinson Franklin Lakes, NJ, USA) and immediately injected into N2-preflushed 12

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mL vials at time 0 and 30 minute after the closure of lids. Air temperature and soil moisture were

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also recorded during each sampling period. Gas samples were analyzed within 24 h on a gas

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chromatograph with an autosampler (Shimadzu AOC-5000 Auto-Injector). Two detectors, a

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flame ionization detector (FID) (Shimadzu GC – 2014, Kyoto, Japan) and an electron capture

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detector (ECD), were fitted with the GC so that CO2, CH4 and N2O can de simultaneously

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measured. N2O was detected by ECD with a detection limit of 0.1 ppm. The N2O fluxes and

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cumulative N2O emissions were calculated using formulas described by Wu et al. (2017).18

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Soil sampling

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We used a 5-cm diameter soil corer to take six to seven soil cores to 15 cm depth after each

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harvesting. Soil cores were separated into 0-7.5 cm and 7.5-15 cm depth fractions. All soil

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samples were sealed in plastic bags, stored in a cooler and transported to the laboratory. In total,

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there are three soil sampling happened in August, September and October for each plot.

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Soil samples were mixed thoroughly and sieved through a 2-mm mesh within 24 h of the

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field sampling and we carefully removed all visible residues and plant roots. Subsamples (~ 20

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g) were then taken immediately and stored at -20 °C for DNA analysis and the remainder of soils



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were stored at 4 °C for other microbial and chemical analyses. A 10-g soil subsample was oven-

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dried at 105 °C for 48 h and weighed for the determination of the water content.

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Chemical and microbial analyses of soil samples

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Soil microbial biomass C and N. Soil microbial biomass C (MBC) and biomass N (MBN) were

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measured using the fumigation-extraction method.56 For each field soil sample, 20-g subsample

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(dry soil equivalent) was fumigated with ethanol-free chloroform for 48 h and then extracted

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with 50 mL of 0.5 M K2SO4 by shaking for 30 min. Another 20-g subsample of nonfumigated

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soil was extracted with 50 mL of 0.5 M K2SO4 after shaking for another 30 min. Soil extractable

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organic C in both fumigated and non-fumigated extracts was determined using a TOC analyzer

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(Shimadzu TOC-5050A, Shimadzu Co., Kyoto, Japan). Soluble inorganic N in the extracts of

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fumigated and non-fumigated soils was quantified on the Lachat flow injection analyzer (Lachat

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Instruments, Milwaukee, WI, USA) after digestion with alkaline persulfate.57 The differences in

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extractable organic C and inorganic N between fumigated and non-fumigated soils were used to

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calculate MBC and MBN using a conversion factor of 0.33 (kEC) and 0.45 (kEN), respectively.56,58

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Soil extractable C and N. The concentration of soluble organic C in non-fumigated soil extracts

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was used to represent soil extractable C. The extractable inorganic N referred to the sum of NH4+

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and NO3- in non-fumigated soil extracts.59

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Soil microbial respiration. An incubation-alkaline method was used to determine soil

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heterotrophic respiration.60 Briefly, a subsample of 20.0g soil (dry soil equivalent) was put into a

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125-mL beaker and adjusted to a moisture level of 60% water holding capacity, and the beaker

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was then placed in a 1-L Mason jar. 5.0 mL of 0.25 M fresh NaOH were added to trap respired

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CO2 in a 50-mL beaker that was suspended in each mason jar.61 All mason jars were incubated at


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25 °C in the dark for 2 weeks. After the first week incubation, NaOH solutions were replaced

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with fresh ones. The respired CO2 from the soil was titrated with 0.125 M HCl to determine the

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soil microbial respiration (SMR). SMR rate was expressed as mg CO2 kg-1 soil d-1 by averaging

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the data across two1-wk incubations.59

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Net soil N mineralization. Potential N mineralization was determined following a 4-wk

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incubation at 25 °C in the dark. After incubation, soil NH4+ and NO3- were extracted with 50 mL

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of 0.5 M K2SO4 following shaking for 30 min. Inorganic N concentrations were also measured

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on the Lachat flow injection analyzer. Net mineralized N (NMN) was then calculated by the

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difference in extractable total inorganic N between un-incubated and incubated soil subsamples.

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Abundances of functional genes responsible for nitrification and denitrification in soil. Fresh soil

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(0.50 g) from final harvest was extracted for nucleic acid using FastDNA SPIN kit (MP Bio,

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Solon, OH, USA) according to the manufacturer’s instructions. Soil DNA quality and size were

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checked by electrophoresis on a 1% agarose gel. The copy numbers of AOA amoA, AOB amoA,

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nirS, nirK and nosZ genes of each soil DNA sample were determined using quantitative real-time

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polymerase chain reaction (PCR) (CFX96 Real-Time PCR Detection System, Bio-Rad,

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Hercules, CA, USA) with the primers given in SI Table S1. The final qPCR reaction volume was

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20 µl, with a of 2 µl of template DNA, 14 µl 1× SsoAdvanced TM SYBR Green Supermix (Bio-

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Rad, Hercules, CA, USA), and 2 µl of each primer. The standard curve for determining the gene

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copy number was developed with the agarose gel-purified PCR products based upon the method

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of Chen et al. (2015).62 All the qPCR reactions followed the conditions were shown in SI Table

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S1.

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Statistical analysis



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We examined results for the entire growing season in 2015 and used the linear mixed model63 to

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test the main effects of temperature, O3 and the interaction of temperature and O3, and whether

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these changed over time. We employed a set of covariance structures including compound

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symmetric model (CS), the first-order autoregressive model [AR (1)], and autoregressive with

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random effect to reduce autocorrelation. The P values for treatments and interaction terms were

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reported based on the covariance structure that minimized Akaike information criterion (AIC)

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and Bayesian information criterion (BIC).63 Data for cumulative N2O emissions and gene copy

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numbers were subjected to the analysis of variance using the mixed model. All statistical

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analyses were performed using the SAS 9.4 (SAS Institute, Inc., Cary, NC, USA). For all tests,

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the significant differences were determined at the 95% probability level.

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RESULTS

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Characteristics of climate change treatments

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During the experiment period, the difference in mean daily temperature between control and

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elevated temperature treatments (T and T+O3) was 3.6 ± 0.07 oC (Table 1 and SI Figure S1). The

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mean daily (12-h) difference in O3 concentration between the control and elevated ozone

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treatments (O3 and T+O3) was 45.6 ± 1.0 ppb (Table 1 and SI Figure S1). The increases in both

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temperature and O3 well met the respective manipulation targets. Meteorological variables (i.e.,

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air humidity, soil moisture and soil temperature) recorded on days of sampling gas indicated

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lower air humidity and higher soil temperature in the warming treatments (T and T+O3)

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compared with the control (CF). No significant differences in soil moisture were found for the

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sampling dates during the experimental study (Table 1).


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Plant biomass

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Warming increased aboveground plant growth at the early stage but this effect faded then after

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(Figure 2a). Across the three sampling dates, warming (T) alone had no significant effect on

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aboveground biomass (Figure 2a). However, it significantly increased root biomass by 22%

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(Figure 2b). In comparison, eO3 had no effect on aboveground biomass (Figure 2a), but tended to

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reduce belowground biomass at the final harvest (-11%) (Figure 2b). There were significant

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effects of the warming × O3 interaction on both aboveground (Figure 2a) and belowground

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biomass (Figure 2b). Warming and eO3 combination (T+O3) aggravated the warming and ozone

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stresses on aboveground plant growth and offset the warming effect on root growth, reducing the

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biomass at the final harvest by 23% and 13%, respectively (Figure 2).

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Soil extractable organic C and N

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Warming had no significant effect on soil extractable organic C in the surface soil (0-7.5 cm),

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but significantly enhanced it in the subsoil (7.5-15 cm) (SI Figure S2 and Table S2). eO3 reduced

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soil extractable organic C (SI Figure S2 and Table S2). Compared to the low ozone plots (CF, T),

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soil extractable organic C under elevated ozone (O3, T+O3) was 12% (P

Contrasting warming and ozone effects on denitrifiers dominate soil

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