Shifts in the composition and activities of denitrifiers dominate CO2

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Shifts in the composition and activities of denitrifiers dominate CO2-stimulation of N2O emissions Yunpeng Qiu, Yu Jiang, Lijin Guo, Lin Zhang, Kent O. Burkey, Richard W. Zobel, S. Chris Reberg-Horton, H. David Shew, and Shuijin Hu Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.9b02983 • Publication Date (Web): 29 Aug 2019 Downloaded from pubs.acs.org on August 30, 2019

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Environmental Science & Technology

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Shifts in the composition and activities of denitrifiers dominate CO2-stimulation of N2O

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emissions

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

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S. Chris Reberg-Horton&, H. David Shew//, Shuijin Hu//,*

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†College

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210095, China

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//Department

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27695, USA

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⊥Institute

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$Institute

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China

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#USDA-ARS,

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&Department

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USA

of Resources and Environmental Sciences, Nanjing Agricultural University, Nanjing

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

of Applied Ecology, Nanjing Agricultural University, Nanjing 210095, China

of Tropical Agriculture and Forestry, Hainan University, Haikou, Hainan, 570228,

Plant Sciences Research Unit, Raleigh, NC 27607, USA

of Crop and Soil Sciences, North Carolina State University, Raleigh, NC 27695,

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1 ACS Paragon Plus Environment


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ABSTRACT

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Elevated atmospheric CO2 (eCO2) often increases soil N2O emissions but the underlying

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mechanisms remain largely unknown. One hypothesis suggests that high N2O emissions may

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stem from increased denitrification induced by CO2-enhancement of plant carbon (C) allocation

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belowground. However, direct evidence illustrating linkages among N2O emissions, plant C

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allocation and denitrifying microbes under eCO2 is still lacking. We examined the impact of

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eCO2 on plant C allocation to roots and their associated arbuscular mycorrhizal fungi (AMF) and

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its subsequent effects on N2O emissions and denitrifying microbes in the presence of two distinct

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N sources, ammonium nitrogen (NH4+- N) and nitrate nitrogen (NO3--N). Our results showed

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that the form of the N inputs dominated the effects of eCO2 on N2O emissions: eCO2

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significantly increased N2O emissions with NO3--N inputs but had no effect with NH4+-N inputs.

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eCO2 increased plant biomass N more with NH4+-N than NO3--N inputs, likely reducing

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microbial access to available N under NH4+-N inputs and/or contributing to higher N2O

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emissions under NO3--N inputs. While eCO2 enhanced root and mycorrhizal N uptake, it also

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increased N2O emissions under NO3--N inputs. Further, eCO2-enhancement of N2O emissions

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under NO3--N inputs concurred with a shift in the soil denitrifier community composition in

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favor of N2O-producing (nirK- and nirS-type) over N2O-consuming (nosZ-type) denitrifiers.

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Together, these results indicate that eCO2 stimulated N2O emissions mainly through altering

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plant N preference in favor of NH4+ over NO3- and thus stimulating soil denitrifiers and their

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activities. These findings suggest that effective management of N sources may mitigate N2O

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emissions by negating eCO2-stimulation of soil denitrifying microbes and their activities.

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TOC/Abstract art

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INTRODUCTION

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Carbon dioxide (CO2) and nitrous oxide (N2O) in the atmosphere are two major greenhouse

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gases, accounting for ~65% and ~7% warming potential, respectively.1,2 Experimental studies

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over the last three decades showed that elevated CO2 (eCO2) in the atmosphere often enhances

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soil N2O emissions,3-7 raising the possibility of amplified N2O emissions under future

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atmospheric conditions. The mechanisms that underlie eCO2-stimulation of soil N2O emissions

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are still unclear, although higher denitrification induced by CO2-enhancement of plant C

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allocation belowground and plant water use efficiency have been proposed as the primary

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mechanisms.6,8,9 However, eCO2-enhancement of belowground C allocation, primarily through

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plant roots and their associated microbes such as mycorrhizal fungi, likely increases plant N

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uptake,10,11 potentially offsetting the CO2 effect on N2O emissions. Yet, no experiment has so far

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been conducted to explore the linkages among plant roots and mycorrhizal fungi, denitrifying

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microbes, and soil N2O emissions as influenced by atmospheric CO2 concentration.

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CO2-enahncement of AMF and plant roots may have contrasting effects on N cycling and

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N2O emissions. Atmospheric CO2 enrichment generally increases plant growth12-14 and

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photosynthate allocation to roots and mycorrhizae.10,15 Arbuscular mycorrhizal fungi (AMF) are

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ubiquitous, form associations with roots of approximately 80% of land plant species and obtain

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C from their host plants in return for mineral nutrients.16,17 On one hand, more root exudates and

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labile C under eCO2 may stimulate denitrification as high C availability likely enhances

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denitrifiers and N2O emissions.6,8,9 On the other hand, large root systems and AMF hyphae likely

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increase plant N uptake and reduce N substrate for N2O production.18-21 However, whether this

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reduction by CO2-enhancement of roots and AMF can quantitatively offset the enhancement of


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N2O emissions under eCO2 remains unexplored.

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Denitrifying microbes are highly diverse, and consisted of ca. 5% of all soil microbes,

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including bacteria and fungi.6,22 Therefore, it is a tremendous challenge (if not impossible) to

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characterize the community structure of denitrifying microbes at this stage.23,24 Fortunately, the

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function genes encoding nitrite reductase (nirK and nirS) and nitrous oxide reductase (nosZ)

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have been well characterized and widely used to describe the abundance and diversity of

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denitrifiers.23-25 Genes nirK and nirS encode two structurally different but functionally

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equivalent enzymes: the copper and cytochrome cd1-nitrite reductase, respectively, that produce

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N2O from the reduction of NO2- to NO.26-28 These two enzymes are thought to be mutually

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exclusive among denitrifiers.29,30 Studies have shown both reductases are functionally redundant,

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as one nir gene in a denitrifying microbe can be eliminated and replaced by the other.31 In

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contrast, the nosZ gene encodes nitrous oxide reductase that convert N2O to N2, that is N2O

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consumption.32,33 Therefore, a high ratio of (nirK+nirS)/nosZ gene copies is usually considered

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to present high N2O production and/or low N2O consumption. Conversely, a low ratio indicates

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either low N2O production, high N2O reduction or both.24 Also, studies have shown that nosZ

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occurs more frequently with nirS than with nirK, indicating that nirK-type denitrifiers contribute

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more to N2O emissions.30,34 The abundances and relative composition of nirK, nirS and nosZ

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have been widely used as marker genes for denitrifying microbes to characterize the dynamics of

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denitrifier communities as influenced by diverse perturbations (e.g. climate warming and N

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deposition).35,36

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CO2-induced alterations in the relative availability between labile C and mineral N may

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critically impact the community composition and activities of denitrifiers. eCO2 often reduces

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plant stomatal conductance and density, and evapotranspiration.37,38 While this enhances plant



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water use efficiency and soil moisture, it may reduce NO3--N uptake as NO3--N often moves to

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plant roots and enters plants with water movement.6,39 Emerging evidence suggests that eCO2

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may promote the preference of C3 plants for NH4+ over NO3- uptake.40-42 A meta-analysis of

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experiments with more than 58 plant species by Cheng et al. (2012)43 also showed that although

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eCO2 reduced soil NH4+, it increased soil NO3-. In addition, Wu et al. (2017)6 have recently

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showed that the form of N inputs dominates the eCO2 effect on soil N2O emissions. Using in-

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field 15N tracing method, Moser et al. (2018)7 also showed that the oxidation of organic N to

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nitrite (NO2-) under eCO2 triggered the enhanced N2O from a temperate grassland at the Giessen

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FACE site, Germany. They further suggested that enhanced activity of nitrite reductase encoded

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by nirS denitrifiers may contribute to the increased N2O emissions, although the nirS abundance

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was not quantified. Together, these results suggest that CO2-enhancement of denitrification may

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control eCO2 effect on N2O emissions, but direct linkages among eCO2, the denitrifiers and N2O

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emissions remain elusive.

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We conducted a microcosm experiment to assess the impact of eCO2 on soil denitrifiers

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and N2O emissions in the presence of AMF alone or both wheat (Triticum aestivum L.) roots and

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AMF. We hypothesized that 1) eCO2 enhances soil N2O emissions via influencing plant N

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preference, which in turn affects denitrifier composition, and 2) promotion of N2O emissions due

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to the CO2-induced change in plant N preference will be quantitatively larger than the

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suppression of N2O emissions resulting from CO2-enhanced plant N uptake via roots and AMF,

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leading to a net CO2-stimulation of N2O emissions.

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MATERIAS AND METHODS

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The experimental design and treatments

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The experiment was conducted in the USDA-ARS Plant Science Research CO2 facility at North

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Carolina State University (Raleigh, NC, USA). The facility consisted of eight continuously

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stirred tank reactors (CSTR) for the exposure of plants to different concentrations of gases (CO2

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in this study). Each CSTR is a cylindrical chamber covered with Teflon and measured 1.2 m in

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diameter by 1.4 m tall.6,43,44 Compressed CO2 was mixed with air and dispensed to chambers 24

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hours daily using rotameters to maintain CO2 concentrations at a target level. To monitor CO2

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concentrations, subsamples of the chamber air were measured using infrared analyzers (model

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6252, LiCor Inc., Lincolin, NE USA) every two minutes per chamber (Figure S1).

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The experiment was a split-split-split plot design with four blocks (replicates) and each

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block consisted of two adjacent CSTR chambers.6 The whole-plot treatments were atmospheric

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CO2 levels (ambient CO2: 400 µmol mol-1 vs. elevated CO2: 680 µmol mol-1). The split-plot

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treatments were two N species (NH4+-N vs. NO3--N), which were further split into three

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mycorrhizal treatments. The two CO2 levels were chosen because the CO2 concentration is

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projected to increase from ca. 400 µmol mol-1 in 2013 to ca. 700 µmol CO2 mol-1 by the end of

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this century.45,46

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There were two plexi-glass microcosms in each CSTR chamber: one was fertilized with

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(NH4)2SO4 and the other with KNO3. Each microcosm was divided into six compartments with

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each compartment measuring 13×14×15cm (width × depth × height) (Figure S2).47 Three

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compartments in a row were designated as HOST compartments (containing host plants and AM

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fungi) and the other three adjacent compartments were designated as TEST compartments to

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determine N2O emission (Figure S2). The HOST and TEST compartments were separated by a

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replaceable 0.45, 20 µm or 1.6 mm mesh (Tetko/Sefar mesh, Sefar America, NY) that controlled



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the penetration of plant roots or AM fungal hyphae into the TEST compartments (Figure S2).47,48

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Three mycorrhizal treatments were therefore formed: The no-AMF control (CK) (0.45 µm) with

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neither AM fungal hyphal nor root penetration, the AMF treatment (20 µm) with AMF only

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growing through, and the AMF+Root treatment (1.6 mm) allowing penetration of both AMF

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hyphae and plant roots. The mesh (0.45 mm) we employed also allows water, N and soluble C

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produced by plant roots to move back and forth between HOST and TEST compartments for all

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the no-root and no-AMF treatments. The AM fungal species mixture used for the experiment

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was trap-cultured from an agricultural soil, collected from the Center for Environmental Farming

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Systems, NC, and was then pot-cultured to increase fungal biomass and spore density in the

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greenhouse. Twelve AM fungal species were identified and characterized according to the

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International Culture Collection of (Vesicular) Arbuscular Mycorrhizal Fungi (INVAM). AMF

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inoculum consisted of culture media containing spores, hyphae, and colonized root pieces.43,47

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AMF inoculation and wheat planting

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Each compartment of the microcosm was filled with a 3.3 kg sandy loam soil collected from the

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Lake Wheeler Research Farm of North Carolina State University, Raleigh, NC, USA (35° 43' N,

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78° 40' W; elevation 120 m). The soil is an Appling sandy loam (fine, kaolinitic, thermic Typic

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Kanhapludult), well drained with a pH of 5.5, and contained 17 g C kg-1 and 1.5 g N kg-1 soil

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prior to the experiment. The HOST compartments were inoculated with 150 g inoculum of a

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mixture of AMF species. Twenty seeds of wheat were sown into each HOST compartment and

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then thinned after emergence to ensure the uniform density. Plants received CO2 fumigation

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immediately following seed germination. The wheat plants were grown in ambient light and

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temperature conditions. To alleviate N limitation of plant growth, 20 mg N kg-1 soil (equivalent


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to 40 kg N ha-1 soil) as NH4NO3 was added to both HOST and TEST compartments at the 4th

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week after germination. HOST compartments with plants were watered with deionized water

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regularly. TEST compartments were also watered but less frequently to maintain soil moisture

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(visually checked every other day) so that AM fungal hyphae and plant roots could grow into the

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TEST soils.

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N2O flux measurement

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At the 8th and 12th week of plant growth, each TEST compartment was watered with fertilizer

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solutions containing NH4+-N as (NH4) 2SO4 or NO3--N as KNO3 at an equivalent rate of 40 kg N

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ha-1 soil, respectively. Equivalent K2SO4 was applied to balance K+. Each TEST compartment

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was watered to reach ca. 80% water-filled pore space (WFPS) in the late afternoon to mimic a

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rain event and ensure an environment conducive for denitrification. Measurement of N2O

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emissions was conducted in the following day when we sampled the N2O once or twice. At the

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8th week of plant growth, N2O emissions were sampled starting at 12, 24, 48, 72, 96 and 120 h

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after fertilizer-N addition, resulting in six measurements of N2O fluxes. At the 12th week of plant

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growth, N2O emissions were sampled starting at 12, 24, 36, 48, 72, 96, 120 until 312 h after

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fertilizer-N addition, resulting in sixteen measurements of N2O fluxes. Because of high

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evapotranspiration induced by the air-circulation system, soil moisture often decreased to ca. 60-

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70% WFPS by the sampling time (i.e., 24 hours after the watering). After N2O sampling in the

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evening, water was again added so that the soil moisture reached to ca. 80% WFPS in the

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evening prior to N2O sampling in the following day.

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N2O fluxes were determined using static chamber and gas chromatography techniques. N2O flux measurements were taken in all TEST compartments using a modified static chamber.



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The chambers were closed with a lid, and fitted with a rubber septum to allow gas sampling via

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syringes. Two gas samples were collected from each static sampling chamber (Figure S2) at 0

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and 30 min after the chamber was closed. We assumed a linear rate of N2O accumulation in the

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chambers over a short period based on previous researches.6,36,49,50 Each sample (5 mL) was

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taken with 20-mL PE syringes (Becton Dickinson, Franklin Lakes, NJ, USA) and immediately

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injected into N2-preflushed 12-mL vial. After each gas sampling, the lid was removed and static

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chambers were kept open. Gas samples were analyzed within 24 h on a gas chromatograph fitted

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with an electron capture detector (ECD) and a flame ionization detector (FID) (Shimadzu GC–

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2014, Kyoto, Japan) with an autosampler (Shimadzu AOC-5000 Auto-Injector). The N2O fluxes

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were calculated using the formulas by Wu et al. (2017)6 and Qiu et al. (2018).36 Mean N2O

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emissions were calculated by averaging the fluxes during each sampling period. At the time of

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gas collection, air temperature and soil WFPS were recorded. We employed a Campbell

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Scientific HS2 Hydrosense II probe (Campbell Scientific, Logan, UT, USA) to measure soil

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volume water contents (VWC, %) to 12 cm depth and then converted the VWC into the WFPS.

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Water addition was terminated when N2O emission was not detectable.

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It is worthy to note that although both nitrification and denitrification produce N2O,

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optimum N2O emissions from denitrification occur in the range of 70-80% water-filled pore

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space (WFPS).51-53 Our experiment was thus designed to target N2O from denitrification, and

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N2O emissions from nitrification were not considered with frequent irrigation of the soil which

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may have well inhibited nitrification.52

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Plant harvest, quantification of mycorrhizal colonization of roots, and plant C and N

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analyses


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Plants were grown for 15 weeks. Plant shoots were cut at the soil surface. Roots were carefully

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separated from the soil and washed thoroughly with tap water. Root colonization by mycorrhizal

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fungi was measured using the gridline-intersect method.54 Subsamples of thoroughly washed

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roots (cut into about 1 cm in length) were cleared in 10% (w/v) KOH, acidified in 1% (v/v) HCl

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for 2 hours, and then stained with acidic glycerol-trypan blue solution.55 The stained roots were

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then spread on a Petri dish with gridlines and examined for infection using a dissecting

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microscope at ×40 magnification. Results obtained were expressed as percentage root length

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colonized (PRLC).47

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Plant shoots and the remaining roots were oven-dried (65C) and weighed. The oven-

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dried shoots were finely ground to powder, and their total C and N concentrations were

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determined using CHN elemental analyzer (Cara Erba and model 2400, Perkin Elmer Co.,

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Norwalk, CT, USA).

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Soil sampling, and soil and microbial analyses

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Soil samples from TEST compartments were taken after plants were harvested, sieved through 2

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mm mesh and stored in 4C refrigerator for analysis. Microbial biomass carbon (MBC) and

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nitrogen (MBN) were determined using a fumigation-extraction method.56 Briefly, a subsample

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of 20.0 g soil (dry soil equivalent) was fumigated with ethanol-free chloroform for 48 h and then

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extracted with 50 mL of 0.5 M K2SO4 by shaking for 30 min. Another 20.0 g subsample of non-

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

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

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TOC analyzer (Shimadzu TOC-5050A, Shimadzu Co., Kyoto, Japan). Soil extractable inorganic

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N referred to the sum of NH4+-N and NO3--N in the extracts of non-fumigated soils. MBN was



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determined following alkaline persulfate oxidation of the fumigated and non-fumigated K2SO4

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extracts.57 The concentration of NO3- and NH4+ was quantified with a Lachat flow injection

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analyzer (Lachat Instruments, Milwaukee, WI, USA). The differences in extractable organic C

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and inorganic N between fumigated and non-fumigated soils were used to calculate MBC and

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

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Quantification of gene copies of functional groups of denitrifiers

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To investigate how eCO2, N forms, and AMF or plant roots affect the communities of

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denitrifying microbes, copy numbers of key genes involved in N2O production (nirK and nirS)

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and N2O consumption (nosZ) were quantified. The ratio of the nirK/nirS or (nirK+nirS)/nosZ

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gene copies has been identified as a useful indicator of the relative composition of the

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denitrifying community and has also been shown to relate to soil N2O production

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capacity.19,24,25,30

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Microbial DNA in soil was extracted from 0.50 g of the fresh field soil prior to the

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experiment and soil samples in the TEST compartments using a FastDNA SPIN kit (MP Bio,

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Solon, OH, USA). The DNA quality and size were checked by electrophoresis on a 1% agarose

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gel. The quantity of the DNA extractions was determined with a nanodrop spectrophotometer

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(Thero Scientific, Wilmington, DE, USA). The copy numbers of nirS, nirK, and nosZ genes

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were determined on each soil DNA sample using quantitative real-time polymerase chain

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reaction (PCR) (CFX96 Real-Time PCR Detection System, Bio-Rad, Hercules, CA, USA). The

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primers for the three genes in the soils were given in Table S1, which were considered as the

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surrogates of the abundances of denitrifying microbes. Each reaction system was performed in a

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20 µL volume involving 14 µL 1×SsoAdvancedTM SYBR Green Supermix (Bio-Rad, Hercules,


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CA, USA), 2 µL of template DNA, and 2 µL of each primer. The standard curve for determining

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

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method of Chen et al. (2015)59 and Qiu et al. (2018)36. All the qPCR reactions followed the

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conditions were shown in Table S1.

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Statistical analyses

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Data were subjected to analysis of variance (ANOVA) using the Mixed Model procedure (Proc

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Mixed) of SAS 9.4. All the data sets of this experiment (split-split-plot design) were analyzed

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using linear mixed-effects model, which tested the effects of CO2 (whole-plot), N form (split-

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plot), mycorrhizae (split-split-plot) and their interactions on all the variables. CO2, N form,

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mycorrhizae and their interactions were fixed effects. The block, whole-plot, split-plot, and split-

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split-plot were random effects. Differences among treatment means were tested post-ANOVA

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using the Tukey-Kramer multiple comparison test.

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RESULTS

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Plant biomass and shoot biomass N

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Compared to the ambient CO2, eCO2 significantly stimulated plant growth (Table 1 and Figure

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1), increasing shoot biomass by 24% (P

Shifts in the composition and activities of denitrifiers dominate CO2

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