Combined Flux Chamber and Genomics Approach ... - ACS Publications

Article pubs.acs.org/est

Combined Flux Chamber and Genomics Approach Links Nitrous Acid Emissions to Ammonia Oxidizing Bacteria and Archaea in Urban and Agricultural Soil Nicole K. Scharko,†,⊥ Ursel M. E. Schütte,‡,⊥ Andrew E. Berke,†,∇ Lauren Banina,† Hannah R. Peel,† Melissa A. Donaldson,† Chris Hemmerich,§ Jeffrey R. White,†,‡ and Jonathan D. Raff*,†,∥ †

School of Public and Environmental Affairs, Indiana University, Bloomington, Indiana 47405-2204, United States Integrated Program in the Environment, Indiana University, Bloomington, Indiana 47405-2204, United States § Center for Genomics and Bioinformatics, Indiana University, Bloomington, Indiana 47405-7005, United States ∥ Department of Chemistry, Indiana University, Bloomington, Indiana 47405-7102, United States

Environ. Sci. Technol. Downloaded from pubs.acs.org by UNIV OF NEBRASKA-LINCOLN on 08/28/15. For personal use only.

‡

S Supporting Information *

ABSTRACT: Nitrous acid (HONO) is a photochemical source of hydroxyl radical and nitric oxide in the atmosphere that stems from abiotic and biogenic processes, including the activity of ammonia-oxidizing soil microbes. HONO fluxes were measured from agricultural and urban soil in mesocosm studies aimed at characterizing biogenic sources and linking them to indigenous microbial consortia. Fluxes of HONO from agricultural and urban soil were suppressed by addition of a nitrification inhibitor and enhanced by amendment with ammonium (NH4+), with peaks at 19 and 8 ng m−2 s−1, respectively. In addition, both agricultural and urban soils were observed to convert 15NH4+ to HO15NO. Genomic surveys of soil samples revealed that 1.5−6% of total expressed 16S rRNA sequences detected belonged to known ammonia oxidizing bacteria and archaea. Peak fluxes of HONO were directly related to the abundance of ammonia-oxidizer sequences, which in turn depended on soil pH. Peak HONO fluxes under fertilized conditions are comparable in magnitude to fluxes reported during field campaigns. The results suggest that biogenic HONO emissions will be important in soil environments that exhibit high nitrification rates (e.g., agricultural soil) although the widespread occurrence of ammonia oxidizers implies that biogenic HONO emissions are also possible in the urban and remote environment.

■

INTRODUCTION

It is well-known that certain microbes present in soil, snow, and water are capable of oxidizing ammonium (NH4+) to nitrate (NO3−) through nitrification.18−20 During this aerobic process, ammonia-oxidizing bacteria (AOB) convert NH4+ into nitrite (NO2¯), while nitrite-oxidizing bacteria (NOB) convert NO2¯ into NO3−. In addition to AOB, the globally distributed Thaumarchaeota (of the domain Archaea)21−23 are also ammonia-oxidizers that have been shown to produce NO2− in laboratory culture studies. 21−26 In some terrestrial ecosystems ammonia-oxidizing archaea (AOA) are more numerous than AOB,21,27 implying that they may be significant contributors to soil NO2− fluxes under certain conditions.21,25,27 A fraction of the soil NO2− formed will be chemically reduced on soil surfaces and contribute to NO and N2O emissions to the atmosphere,28−30 while some will be

Nitrous acid (HONO) is an important photochemical source of hydroxyl radical (OH) and nitric oxide, which play a role in initiating and propagating the chain reactions leading to ozone and aerosol formation in the polluted atmospheric boundary layer.1 Since the first detection of HONO in the lower atmosphere,2,3 efforts have focused on understanding the sources of HONO. Studies carried out in both urban and rural areas have observed vertical gradients of HONO (high concentrations near the ground), suggesting that HONO is formed by processes occurring at ground level.4−9 Most of the mechanisms proposed are abiotic and involve heterogeneous or photochemical reactions of nitrogen oxides (e.g., NO, NO2, HNO3, and NO3¯) on ground surfaces,10−14 although recent evidence suggests there are potentially significant biogenic sources of HONO in snow and soil that could contribute to HONO emissions throughout the day.15−17 In this study, we combine laboratory mesocosm measurements of HONO fluxes with genomic analyses of mesocosm soil to show that biogenic emissions of HONO are related to the abundance of nitrifying microbes indigenous to agricultural and urban soil. © XXXX American Chemical Society

Special Issue: Ron Hites Tribute Received: February 15, 2015 Revised: July 21, 2015 Accepted: July 23, 2015

A

DOI: 10.1021/acs.est.5b00838 Environ. Sci. Technol. XXXX, XXX, XXX−XXX

Article


Environmental Science & Technology

was used for HONO flux measurements. The second sampling location was Bryan Park, a municipal park located in a residential neighborhood near downtown Bloomington, IN. Bloomington is a city with population of over 82 500 that is located in Monroe County, IN. Samples were collected from two different sites on the park grounds (39.16 N, −86.53 W). The Park-1 sample was collected from the center of the park, near the edge of a lawn area (dominated by Timothy grass, Phleum pretense) that receives minimal foot traffic. The Park-2 sample was collected from mostly bare soil located under the canopy of a Norway spruce (Picea abies) that was growing adjacent to a busy pedestrian walkway. The Park-3 sample was collected from the identical location as the Park-1 sample, but at a later date after the Park-1 sample had been depleted. Management of park grounds is limited to the occasional mowing; no fertilizer or pesticides were applied. Prior to experiments, fresh soil was distributed on a tray and allowed to air-dry overnight. Samples were then sieved with a 10 mesh (2 mm) soil sieve and stored separately in plastic containers at ambient temperature (22 °C) in the dark and used within 2 weeks of collection. Soil pH was measured in a 1:5 soil/water (v/v) suspension using a Thermo-Scientific Orion Star A211 and Ross Ultra pH/ATC triode.40 Determination of NH4+ and NO3− concentrations in soil prior to use was performed at the University of Minnesota (St. Paul, MN) using established methods.41 Flux Chamber Measurements. Soil samples were uniformly distributed on 8.2 × 8.2 × 0.2 cm3 aluminum trays, and sealed in 1 L flux chambers made of DuPont Tedlar (polyvinyl fluoride) and equipped with inlet and outlet ports and valves. The aluminum trays and all exposed metal surfaces present in the experimental system were coated with an inert perfluorinated polymer (Fluoropel PFC 801A, Cytonix Corp.); Teflon tubing and fittings were used throughout. Perfluorinated materials were used to reduce the adsorption and reactivity of oxides of nitrogen with laboratory surfaces that could interfere with measurements. All soil samples in the flux chambers were kept in the dark at ambient temperature (22 °C) during measurements and incubation. Three sets of experiments were carried out, the results of which are displayed in Figure 1. Each set of experiments was comprised of up to four chambers operated in parallel under the same conditions. One chamber (labeled “empty chamber” in Figure 1) contained no soil, while the other three chambers contained an equivalent mass of soil that contained one of the following treatments: (i) deionized water (18 MΩ cm; Milli-Q Integral); (ii) an aqueous solution of (NH4)2SO4 (45 mM, Macron Fine chemicals, AR ACS granular); or (iii) an aqueous solution of (NH4)2SO4 (45 mM) and nitrapyrin (13 mM, 2chloro-6-(trichloromethyl) pyridine ≥98%, Sigma-Aldrich), in the case of the Ag-2 and Park-1 soil. The amount of soil added to flux chamber (23 g for Ag-1 and -2; 16 g for Park-1, -2, and -3) was chosen based on the amount of soil needed to completely cover the bottom of the tray and to the full depth of 0.2 cm. We confirmed in separate tests that 16 and 23 g of soil produced no significant difference in peak HONO emission flux. Soils were amended through dropwise addition (1.7 mL total volume) of one of three treatments described above to a 50 g batch of soil using a micropipette dispenser. The wetted soil was then mixed (hand shaken for 1 min) until the soil appeared homogeneous. The (NH4)2SO4 amendments increased the NH4+ concentrations by 55 μg g−1 relative to the amount present at time of collection; the final concentration of

released from soil as HONO.16 The latter process will depend on soil water content,17 relative humidity,31 soil water pH,32,33 the deposition of strong acids capable of displacing HONO,34,35 and the surface acidity of minerals present in the soil.36 Although NO 2 − is well-known to be an important intermediate in the terrestrial nitrogen cycle, evidence that it may be a significant source of HONO to the atmosphere has only recently emerged. Kubota and Arimi provided the earliest evidence that HONO emitted from soil could have a biological origin.37 Amoroso et al. attributed measured HONO fluxes below the surface of snowpack at Ny-Ålesund (Norway) to microbial activity after finding that the 17O and 15N isotopic signature of NO2− and NO3− extracted from the snow was characteristic of fractionation by biological processes. 15 Maljanen et al. subsequently showed that the intrinsic HONO emissions from acidic soil collected from drained peatlands were higher than those from pristine peatlands with a high water table, suggesting that aerobic nitrification contributed to HONO emissions in the former biome.38 Oswald et al. showed that HONO and NO fluxes from soil collected from arid and arable regions of the globe were comparable in magnitude, and observed release of NO and HONO when an AOB culture (Nitrosomonas europaea) was added to a model soil substrate.16 The same group used a tracer method based on an azo dye derivatization technique and high performance liquid chromatography−mass spectrometry to identify HO15NO emitted from soil treated with 15N-labeled urea.39 While recent efforts have provided convincing evidence for the existence of a biogenic source of HONO, a full microbiological characterization of the soil samples exhibiting HONO production is still lacking. We hypothesize that the type of NH3 oxidizing microbes present and their relative abundance will be key factors in determining the HONO emissions from soil. To test this, we carried out flux chamber experiments on soil samples collected from agricultural and urban sites in southern Indiana to identify soil that emitted HONO into the gas phase. The biogenic nature of these HONO emissions was probed in a series of experiments where NH4+ or a nitrification inhibitor were added, and from stable isotope tracer experiments aimed at demonstrating 15NH4+ → HO15NO conversion in soil. The 16S rRNA genes and expressed rRNA from the samples were sequenced to provide information on the microbial community composition and activity. To our knowledge, this represents the first genomic survey of the microbial consortia indigenous to agricultural and urban soil shown unambiguously to emit HONO. These observations are combined with the current understanding of the ecology of AOA and AOB to provide insights into the environmental conditions and biome types where biogenic HONO emissions are important.

■

EXPERIMENTAL SECTION Sampling Sites. Samples were collected by pushing aside surface vegetation and recovering the uppermost soil layer (down to 5 cm) at two different locations in Southern Indiana; site and soil characteristics are listed in Table S1. The first location is an agricultural field located in rural Bartholomew County, IN (39.17 N, −85.89 W). Samples were collected from the same site in a soybean (Glycine max) field on two separate occasions (May 2013 and July 2014); these samples are labeled Ag-1 and Ag-2, respectively. The Ag-1 soil was used to develop the methods used for the genomic survey, while the Ag-2 soil B


Article



ppb of HONO) diffusing through the chamber walls. Thus, measurements of HONO in the “empty chamber” experiments represent the background levels of HONO expected to be present in all of the treatments on a given day. This defines the CEAS limit of quantitation for HONO fluxes at 0.5−1 ng m−2 s−1. The 2σ statistical error in flux measurements is on average ±0.7 ng m−2 s−1, and is associated with the spectral fits. Chemical Ionization Mass Spectrometry Measurements. Chemical ionization mass spectrometry (CIMS) was used to measure HONO or HO15NO emitted from soil that was amended with either unlabeled or 15N-labeled ammonium sulfate. This allowed us to follow microbial oxidation of 15NH4+ to HO15NO directly and in real-time. Substrates were amended with 1.7 mL of a 45 mM aqueous solution of (14NH4)2SO4 or (15NH4)2SO4 (99 atom % 15N, Cambridge Isotope Laboratories Inc.) using the procedure described above and placed inside separate flux chambers; HO 14 NO and HO 15NO were monitored for an hour each day for 2 consecutive days at a flow rate of 2100 cm3 min−1 and RH of 50%. Under these conditions, the residence time of headspace air was 30 s. Soil chambers were sealed in the dark and in an atmosphere of humid air in between measurements. The experiment was repeated for two soil samples (Ag-2 and Park-3) and a sterile blank consisting of kaolinite (Fluka Analytical). The CIMS instrument uses SF5− as the reagent ion to selectively detect HONO as [NO2]− at m/z 46, whereas HO15NO is detected as [15NO2]− at m/z 47. Operation of the CIMS instrument and its calibration have been described previously.44 For an integration time of 0.2 s, the detection limit for HONO (m/z 46) was 660 ppt, with a standard deviation of 65 ppt for a 10 min measurement; the detection limit for HO15NO (m/z 47) was 80 ppt, with a standard deviation of 13 ppt for a 10 min measurement. Microbial Community Characterization. The bacterial and archaeal community composition in soil samples was assessed by sequencing extracted 16S rRNA genes. This gene is common to all bacteria and archaea and its sequence provides information on community composition, although it does not discriminate which members are biologically active. To identify the active members, RNA was also extracted and sequenced. For both DNA and RNA we targeted the 16S rRNA gene using the primers 515f and 806r as described by the Earth Microbiome Project.45,46 Thus, two sets of sequences are presented in this study: The 16S rRNA gene sequences derived from extracted DNA, and the expressed 16S rRNA gene sequences derived from total extracted RNA. To avoid confusion, we will refer to the 16S rRNA genes as the “DNA,” while the expressed rRNA genes will simply be called “RNA.” As the species definition of bacteria and archaea is an area of debate, sequences with 97% similarity are clustered into operational taxonomic units (OTUs) before matching the resulting consensus sequence of each OTU to known sequences in the SILVA database.47 Each OTU is then used as a proxy for representing a microbial species present in a sample, where the relative abundance of an OTU is directly proportional to the abundance of the microbial taxa that this OTU describes.48 Additional details, including procedures for genomic DNA and RNA extraction from soil, sequencing, and data reduction, are provided in the Supporting Information.

Figure 1. Flux chamber measurements showing nitrous acid (HONO) emissions from soil samples collected from an agricultural field and at two different sites in an urban park located in Bloomington, IN. Each soil sample was amended with the indicated treatments at day 0 and sampled for a period of 1 h each day. Error bars represent uncertainty of the least-squares fit of reference spectra to the measured spectra. For each sample, the 3−4 treatments shown were run in parallel.

nitrapyrin in the inhibited soils was 102 μg g−1. Separate tests confirmed that the pH of the soil did not change immediately after adding the various amendments. The resulting soil mixtures had a water content of between 21 and 23%, as determined gravimetrically from each sample before and after drying at 125 °C for 24 h. HONO Flux Measurements. Nitrous acid fluxes were measured using a home-built cavity enhanced absorption spectrometer (CEAS) that has been described previously (see also SI).42,43 For each measurement, the inlet and outlet valves of the chamber were opened to a flow of zero air [500 cm3 min−1, 1 atm, at 50% relative humidity (RH)] that carried gases emitted into the headspace of the chamber to the CEAS cavity for HONO detection. The RH was controlled by flowing air through a glass frit submerged in deionized water, followed by dilution with dry air in a 3 L reservoir equipped with a relative humidity gauge and temperature probe (Vaisala, HMT130). The residence time of air in the chamber headspace was ∼2 min. The concentration of HONO flowing through the chamber was derived by averaging 180 spectra (spectral integration time of 20 s) collected during a 1 h measurement period. Each soil chamber was kept sealed in the dark and in an atmosphere of humid air for between 1−24 h in between measurements to minimize soil moisture loss. Each chamber was monitored in the dark for 1 h per day and for 5 consecutive days, except for Park-2 samples, which were monitored for 9 days. Ultrapure air was supplied to experiments from a zero air generator (Parker-Balston). Nitrous acid fluxes were determined using, HONO Flux = C × F/A, where HONO Flux has units of ng m−2 s−1, C is the HONO concentration (in ng m−3) measured by the CEAS, F is the flow of air through the chamber and CEAS (m3 s−1), and A is the geometric surface area (m2) covered by the soil sample in the chamber. The sensitivity of the CEAS is limited by the amount of background HONO present in the chamber, which varies over the course of the experiments. Measurements using an empty chamber were collected in parallel to determine daily HONO background levels (Figure 1; labeled “empty chamber”). The HONO measured in the empty chamber experiments stems from residual HONO outgassing from the CEAS surfaces, tubing, or laboratory air (indoor levels of 1−2

■

RESULTS HONO Fluxes from Agricultural and Urban Soil. Addition of deionized water to Ag-2 and Park-1 samples to a C


Article


Environmental Science & Technology soil water content of 22% (w/w) did not yield a detectable increase in HONO flux above background levels detected in empty chambers (Figure 1). In contrast, a sharp increase in HONO emission was observed within an hour of NH4+ addition to soil, with fluxes peaking 24 h after amendment at 19 and 8 ng m−2 s−1 for Ag-2 and Park-1 samples, respectively. The slight increase in HONO flux for Park-1 samples measured from 72 to 96 h is due to higher background levels of HONO. A sharp rise in HONO flux after 1 h was not observed following addition of NH4+ to Park-2 soil. Instead, the HONO fluxes for this soil gradually increased and only peaked after 96 h. Interestingly, the peak HONO fluxes observed from the Park-2 soil treated with a NH4+ solution and pure water are nearly identical within experimental error. The peak HONO fluxes in both treatments are ∼5 times higher than what was measured in the empty chamber on this day, which suggests that the peak is due to emission from the soil. Using the peak HONO flux as an indicator of the potential for biogenic HONO formation, the results suggest that microbial activity responsible for HONO formation follows the trend, Ag-2 > Park-1 > Park-2. Inhibitor and Stable Isotope Tracer Studies. The connection between HONO fluxes observed in chamber incubation and microbial activity was further investigated with two complementary approaches. First, soil from Ag-2 and Park1 was amended with a mixture of NH4+ and nitrapyrin, which is widely used to inhibit nitrification during injection of anhydrous ammonia in soil prior to crop planting.49−51 Inhibition of nitrification is a strategy to increase NH3 availability to crops and reduce acidification of soil. As shown in Figure 1, HONO fluxes from soil treated with nitrapyrin and NH4+ were comparable to those of soil treated with water. The ability of nitrapyrin to suppress HONO production in Ag-2 and Park-1 samples strongly suggests that HONO is generated during the process of nitrification. In the second approach, a stable isotope tracer was used to quantify biogenic contributions to HONO emissions in Ag-2 and Park-3 soil samples. Soil samples were amended with 15 NH4+ and chemical ionization mass spectrometry (CIMS) was used to distinguish between HONO emitted by microbial activity and that produced by abiotic mechanisms in real-time. Figure 2A shows the concentration of HO15NO in the flux chamber 2 days after soils were amended with an aqueous solution of (15NH4)2SO4. Experiments were initiated by measuring background concentrations of HO15NO in humidified air. At ∼ 10 min, the humidified air was flowed through a chamber containing soil, resulting in an instantaneous increase in the m/z 47 signal, due to the [15NO2]− ion. For Ag-2 soil, the HO15NO concentration initially reached ∼9 ppb, and continued to increase over the course of an hour to ∼12 ppb. This suggests that the production rate for HONO by nitrifiers was greater than the loss rate due to the purging the chamber. For Park-3 soil, HO15NO peaked at 2 ppb, and gradually decreased over the course of an hour. Thus, microbial production rate of HONO in Park-3 soil was lower than the removal rate from the chamber by the purge. The increasing HO15NO emission rate observed for the Ag-2 soil and diminishing emission rate seen for Park-3 indicates the nitrifier community is more active in agricultural soil. Day two concentrations of HO15NO and HO14NO are presented for 15NH4+ isotope tracer experiments (Figure 2B), and for unlabeled 14NH4+ addition experiments (Figure 2C). When 15NH4+ was used, we observed 2−3 times more

Figure 2. (A) Concentration of HO15NO measured as [15NO2]− (m/z 47) by chemical ionization mass spectrometry (CIMS) when flux chambers containing agricultural and urban soil are sampled on the second day after amendment with an aqueous solution of (15NH4)2SO4 (isotopic purity of 99%). The other panels show HONO measured by CIMS when flux chambers containing agricultural and urban soil and a kaolinite blank are sampled on the second day after amendment with an aqueous solution of: (B) (15NH4)2SO4, 99 atom % 15N, or (C) (14NH4)2SO4. The [HO15NO]: [HO14NO] ratio appears above the bars. Measurements by CIMS were taken by sampling a stream of air (50% relative humidity) flowed through the flux chamber.

HO15NO in Ag-2 and Park-3 samples than HO14NO (Figure 2B). The residual HO14NO emitted from Ag-2 soil (Figure 2B) is attributed to microbial processing of the existing NH4+ pool that was present in the soil at the time of sampling (Table S1). For comparison, traces of HONO present during control experiments involving addition of 15NH4+ to sterile kaolinite (a clay mineral identified in these soils via X-ray powder diffraction) 36 were in the form of HO14NO, whereas HO15NO was not measured above the detection limit of 80 ppt. Kaolinite is a Lewis basic alumino-silicate clay mineral with a high surface area that absorbs oxides of nitrogen from ambient air (e.g., during storage or handling).52,53 Thus, the low concentrations of HO14NO still measured when 15NH4+ is added to kaolinite are likely due to outgassing of background HONO from the clay surface. When 14NH4+ is added to the soil (Figure 2C), the majority of the total HONO is in the form of HO14NO, while HO15NO was below the detection limit of 80 ppt. The ratios displayed above the bars in Figure 2C and in the case of the kaolinite blank shown in Figure 2B, were derived using the HO15NO detection limit, leading deviations from the 1:250 ratio predicted by natural abundance of 15N/14N. Genomic Survey of Soil Microbial Communities. Expanding on the finding that the HONO soil emissions are of biogenic origin, we determined the bacterial and archaeal community composition and activity in the soils used for the flux chamber experiments (Figure 1) using barcoded amplicon sequencing of the DNA and RNA. Matching the consensus sequences against known sequences in the SILVA database, it was determined that 2−10% of all the sequences detected in soils from this study were members of the following Noxidizing microbial taxa: Ammonia-oxidizing archaea (AOA, 12 OTUs), ammonia-oxidizing bacteria (AOB, 60 OTUs), and nitrite-oxidizing bacteria (NOB, 52 OTUs); see Tables S2−S5. D


Article



Figure 3. (A) Comparison of the relative abundance of DNA (solid bars) and RNA (hatched bars) gene sequences attributed to ammonia oxidizing archaea (AOA), ammonia oxidizing bacteria (AOB), and nitrite oxidizing bacteria (NOB) in the indicated soil samples. Samples Ag-2, Park-1, and Park-2 were amended with aqueous (NH4)2SO4 and incubated prior to genomic analysis; sample Ag-1 was untreated with ammonium prior to genomic analysis. The other graphs show variation in operational taxonomic units (OTU) across the soil samples describing differences in community composition (DNA) and activity (RNA) for (B) AOA, (C) AOB, and (D) NOB. The definition of each OTU is provided in the Supporting Information.

interactions with the AOB.22 The results show that the relative abundance of AOB in these soils follows the trend, Ag-2 > Park-1 > Park-2, similar to the trend for HONO fluxes shown in Figure 1. It is possible that the lower AOB population observed in Park-2 soil was influenced by the presence of allelopathic plant root exudates (e.g., monoterpenes from the nearby Norway spruce) that have been shown in some studies to inhibit nitrification.60−62 However, as discussed below, we believe a more plausible explanation is that the AOB were inhibited by the low soil pH at this site. Figures 3B−D show the relative proportion of the AOA (Figure 3B), AOB (Figure 3C), and NOB (Figure 3D) members in soils from Ag-2, Park-1, and Park-2. The community composition and activity of Ag-2 and Park-1 samples were similar, but differed significantly from the Noxidizers present in the Park-2 sample. For example, one member of the Nitrosomonadaceae family (OTU 55), comprised 58% of AOB in Park-2, compared to 17% and 20% in the Park1 and Ag-2 samples. OTU 109 matched a known sequence from the South African Gold Mine Group-1 (SAGMCG-1) within Thaumarchaeota and made up 70% of the AOA in Park2, compared to 0.028% and 0.005% in the Park-1 and Ag-2 samples (Figure 3B). These trends were surprising considering Park-1 and -2 were sampled within 120 m of each other, while the Ag-2 sample was collected in Bartholomew County, some 60 km away. Figure 3D shows that there are distinct shifts in the nitrite oxidizers present and those NOB that were active. For example, in sample Park-1 Nitrospira (OTU 30) made up 59% of the nitrite oxidizers present but only 8% of the active nitrite oxidizers. However, a member of Nitrospirales (OTU 111) was present at 4% but made up 21% of the active nitrite

The types of AOA, AOB, and NOB we found are consistent with other studies. With the exception of one OTU, all archaeal OTUs matched known sequences within the Thaumarchaea,54,55 which are AOA; most were within the Soil Crenarchaeotic Group (SCG). A significant portion of OTUs detected in our samples belonged to Nitrosomonadaceae, a family well-known for obtaining its energy by oxidizing ammonia to nitrite.20 The other ammonia oxidizer detected belongs to the genus Nitrosococcus.56 As for NOB, our sequences matched those of well-described nitrifiers in the Nitrospinaceae family,57 Nitrospira, and other members of the order Nitrospirales.58,59 Figure 3A shows the fraction of DNA and RNA sequences belonging to AOA, AOB, and NOB in soil samples from Ag-1, Ag-2, Park-1, and Park-2 sites. The solid bars are the DNA sequences of those community members present that signify the background genetic potential of the soil. The hatched bars are the RNA sequences, representing the abundance of the active members. All samples, with the exception of Ag-1 were those used for the NH4+ amended trials shown in Figure 1. The Ag-1 sample was collected from the same site as Ag-2 and serves as an example of the N-oxidizer community present in soil that was not amended in the laboratory prior to genomic analysis. Interestingly, there is a higher fraction of AOB and NOB in the RNA than in the DNA across all samples, except for the Park-2 sample where the NOB were rare. This suggests that the relative abundance of active AOB and NOB was larger compared to the DNA sequences attributed to AOB and NOB in the soil. In contrast, the percentage of archaeal RNA was lower compared to the percentage of archaeal DNA present in the samples (Figure 3A), which may be due to competitive E


Article


Soil Acidity and HONO Fluxes. The low soil pH measured in Park-2 soil likely explains the low abundance of AOB (Figure 3) and correspondingly low HONO fluxes for this soil (Figure 1). Since the Park-2 site is adjacent to a busy pedestrian pathway, it is possible that animal waste delivered by pet traffic could have contributed to the high NH 4 + concentration, which would lead to acidification and buildup of NO3¯ over time.71,72 Low soil pH limits the availability of NH3, eventually leading to reduced nitrification rates.73,74 While some AOB are capable of ammonia oxidation at low pH if they grow on alkaline microsites within the soil or via intracellular urease activity,74 most cultured representatives show little-to-no activity below pH 6.5.75 For example, plotting relative nitrification rates (RNR) as a function of soil pH from several reported studies76−78 shows that nitrification rates are directly correlated to soil pH (Figure 4A). Lower nitrification

oxidizers. Similarly, a member of Candidatus Entotheonella (OTU 21) made up 42% of the active community but was only present at 6% in the Ag-2 sample.


■

DISCUSSION The following observations strongly suggest that HONO emitted from agricultural and urban soil stems from indigenous populations of aerobic ammonia oxidizing microbes present in the soil: (1) HONO emissions were stimulated through addition of NH4+, but inhibited by addition of a nitrification inhibitor; (2) a high [HO15NO]/[HO14NO] ratio is observed for soil treated with 15NH4+; (3) HONO fluxes are directly proportional to the abundance of active NH3 oxidizers in the soil; (4) the notable absence of genes attributable to denitrifying bacteria confirms that aerobic nitrification is the principal source of biogenic HONO in these samples. The results further highlight that nutrient availability is a key factor in the potential of soil to emit HONO, which is in turn dependent on edaphic conditions. Effect of Soil Moisture on HONO Fluxes. During NH4+ amendment studies for Ag-2 and Park-1 soil (Figure 1), the initial increase in HONO flux within the first hour is likely related to the time it takes for the microbial communities to recover their functional capacity following a period of inactivity under dry soil conditions. The subsequent decline in HONO emissions after 24 h is likely due to decreasing nitrifier activity in response to nutrient starvation induced by low soil moisture levels.16 Over the course of each experiment, the soil water content decreased from 22 ± 1% (w/w) at wet-up, to as low as 9% on the last day of the experiment due to the high flow of air through the chamber during the measurements. As discussed previously, maxima in HONO emission fluxes are expected when soil water content is between 15−30%.16 This represents a condition where ample water is present to support nitrifier activity, yet the water content is low enough to facilitate diffusion of O2 and HONO through the soil. Increased mobility of nutrients is also likely responsible for profiles in HONO emission fluxes observed for the Park-2 samples in Figure 1. Rewetting of the soil appears to have been sufficient to mobilize the high concentrations of NH4+ (24 μg g−1 soil) present in this soil at the time of collection. Amending the Park-2 soil with (NH4)2SO4 only increased the soil NH4+ concentrations by 1%, compared to a 27% increase in the case of the Ag-2 and Park-1 samples. This explains why there was not a noticeable difference in the peak HONO flux when Park-2 soil was treated with water versus when it was treated with an NH4+ solution. The trend in HONO emissions observed in Figure 1 is consistent with the known response of microbes to soil rewetting events, such as those observed during the first rain following a dry period.63−68 Large pulses of NO and N2O during such events have been attributed to the resuscitation of dormant microbial populations as water addition removes diffusion limitations and increases the availability of watersoluble nutrients.28,69,70 Placella and Firestone measured an increase in the quantity of bacterial ammonia monooxygenase (amoA) transcripts within an hour of wet-up and found that despite months of inactivation in dry soil, AOB and some strains of NOB were able to respond rapidly to increased moisture levels by upregulating amoA gene expression to compete for nutrients made available by soil rewetting.66 In these studies, upregulation of nitrifier gene transcripts was accompanied by an increase in soil nitrate concentrations.

Figure 4. (A) Relative nitrification rates in soil (RNR)59−61 and (B) emission fluxes of HONO from soil measured in laboratory incubation studies16,29 are similarly correlated to soil pH. Equations of the linear fits to the data are ln(RNR) = (0.84 ± 0.12) × pH − (6.37 ± 0.72); ln(HONO flux) = (0.75 ± 0.16) × pH − (0.75 ± 0.16), where HONO fluxes are in units of ng m−2 s−1. In the upper graph, net nitrification rates from the indicated studies were normalized to the maximum nitrification rates to allow for comparison between soil sampled from different biomes.

rates in the Park-2 sample would in turn lead to accumulation of NH4+ and lower NO2− levels; this likely explains the low NOB abundance in this soil.58 We also note that addition of 55 μg g−1 of NH4+ to Park-3 soil caused the soil pH to drop by 0.6 pH units over the course of 6 days. Thus, it is possible that declining soil pH may have contributed to decreased HONO fluxes observed in the later stages of the experiments described in Figure 1. To further assess the importance of soil acidity on biogenic HONO emissions from soil, we compared the pH dependent HONO flux data available from this and previously reported laboratory-based soil chamber studies. Figure 4B shows the natural logarithm of HONO fluxes reported from laboratory flux chamber studies plotted against soil pH. Linear regression F


Article



The lower NH3 availability in acidic soils73,74 is expected to affect the abundance of active ammonia-oxidizing bacteria but likely influence AOA less as the archaea have a higher affinity for NH3.24,75,86,87 The time delay required to reach peak HONO flux for Park-2 soil (Figure 1) compared to the rapid response to rewetting seen for Ag-2 and Park-1 soil would also support a role for AOA as a HONO source in these experiments. Placella and Firestone observed that AOA lagged behind AOB in their response to wetting of dry soil collected from an arid grassland, although AOA were able to maintain elevated transcription of amoA genes for a longer period of time.66 Environmental Implications. An important question is whether NH3-oxidizers in soil are capable of producing atmospherically significant fluxes of HONO and whether they contribute to HONO concentrations in ambient air. The biogenic HONO fluxes summarized in Figure 4B are mostly in the range of 0.5−40 ng m−2 s−1, with two agricultural soil samples reported by Oswald et al. exhibiting HONO fluxes around 250 ng m−2 s−1.16 These fluxes can be compared to HONO fluxes measured during recent field studies in urban and rural regions (Table S6). Although large diurnal variability is observed in each study, the ranges of HONO fluxes across all field studies are consistent, falling between a minimum of −24 (net deposition) and a maximum of 61 ng m−2 s−1 (net emission).88−91 While many of the biogenic HONO fluxes derived from laboratory flux chamber experiments are consistent with field observations, it is not possible to determine whether any of the field studies were influenced by biogenic emissions without knowing more about the microbial communities present or the soil properties at these various sites. In general, agricultural soil that is primed by years of fertilization is expected to have a higher density of nitrifying organisms capable of leading to HONO emissions. Biogenic emissions of HONO from soil are potentially important in agricultural areas, where large pulsed emission events are expected following fertilizer application, or when cropland is irrigated or rained on following a dry period. Additionally, the resemblance of the microbial composition in the Park-1 and Ag-2 soil samples suggests that HONO emissions from ammonia-oxidizing activity in soil is a widespread phenomenon that extends to urban soils as well. This is consistent with nitrogen cycling studies focused on urban lawns, forests, and riparian regions, which have shown that these ecosystems have a high capacity to retain nitrogen from atmospheric deposition, runoff, or fertilizer application, and support nitrifier activity.92−97 However, it remains to be seen whether biogenic HONO emissions are important in urban areas with high NOx concentrations, where photochemistry, heterogeneous reactions, and direct emissions from combustion may be the dominant HONO sources.5,98 Lastly, it is possible that microbial activity may be a dominant source of atmospheric HONO in remote areas where ambient NOx levels are typically low. This includes arctic regions where AOA have been shown to outnumber AOB and are major players in determining the nitrification activity of this nitrogen-limited ecosystem.25 Future studies aimed at determining whether biogenic HONO fluxes have an important impact on atmospheric composition will require concerted field, mesocosm, and modeling studies aimed at determining the relative importance of photochemical, thermal, and biogenic sources.

of the data shows that there is a statistically significant correlation between HONO flux and soil pH, with HONO emissions increasing as soil pH rises. The fit to the data is good considering the diverse biomes sampled. The fact that all the data compiled were collected in a controlled laboratory environment and the peak HONO fluxes are reported helps control for temperature and differences in soil water content in the field. The slopes of linear equations used to fit the data in Figure 4B and Figure 4A are remarkably similar, suggesting that the effect of soil pH on nitrification rate and hence NH3 availability exerts an important control on HONO emissions to the atmosphere. In contrast, no relationship was found between HONO emission fluxes and NH4+ concentrations measured in soil samples. This is consistent with previous studies that showed poor correlations between soil NH4+ concentrations and nitrification potential.64 Although two of the highest HONO fluxes observed by Oswald et al. were associated with high NH4+ concentrations, numerous samples did not follow this trend.16 In our study, the Park-2 sample had the lowest HONO emission rate, although the soil contained the highest NH4+ concentration of the sites studied (Table S1). However, the low soil pH in the Park-2 sample was likely responsible for lower abundance of active AOB. Numerous processes will moderate NH4+ and NO2− concentrations in different ways leading to poor correlations between these nutrients. This includes net N mineralization rates, oxidation of nitrite by NOB, and reactions of HONO in the soil (e.g., with carbonates that sequester HONO as nitrite35 or via chemodenitrification79−81). Thus, nitrification rates rather than NH4+ concentrations in soil are likely a better predictor of a soil’s propensity to emit HONO. While soil pH controls nitrification rates, it also plays an important role in determining the speciation of N(III), which includes H2NO2+, HONO, and NO2−. Until recently, it was thought that the partitioning of HONO between soil and air was governed by bulk aqueous chemistry and Henry’s law, which predict that HONO would only be emitted into the atmosphere at low soil pH (e.g., pKa for HONO is ∼3).32 This view is inconsistent with the data compiled in Figure 4B, where HONO emissions tend to be higher under neutral or alkaline soil conditions.9,16,17,38,82 Recently, it was found that amphoteric Al and Fe (hydr)oxides present in soil are capable of protonating NO2− well above the pKa of HONO and provide a way to sustain HONO emissions at near neutral pH.36 The optimal soil for HONO emission will be one that supports a diverse and abundant population of ammonia oxidizers (pH 7− 9) in a soil whose mineral composition is replete with amphoteric Al and Fe-based mineral oxides. Contributions of AOA to HONO Fluxes. The detection of archaeal RNA in all soil samples suggests that AOA may have contributed to HONO soil emissions observed in this study, although their relative contribution remains unclear. The higher abundance of AOB sequences relative to AOA sequences in Ag2 and Park-1 soil (Figure 3A) would suggest that AOB are the dominant source of HONO for these samples. However, AOA may play a more important role in the HONO emissions observed from the Park-2 sample, where AOA comprised a third of the total NH3-oxidizing community. Previous work has established that low soil pH is associated with low bacterial diversity, while archaea do not display a similar dependence on pH.22,83,84 For example, whereas the gene abundance of bacterial amoA genes decreased with lower pH, gene abundance of archaeal amoA genes increased in an opposite trend.21,27,85,86 G


Article


■

gradients during SHARP 2009 in Houston, TX. Atmos. Chem. Phys. 2012, 12, 635−652. (8) VandenBoer, T. C.; Brown, S. S.; Murphy, J. G.; Keene, W. C.; Young, C. J.; Pszenny, A. A. P.; Kim, S.; Warneke, C.; de Gouw, J. A.; Maben, J. R.; Wagner, N. L.; Riedel, T. P.; Thornton, J. A.; Wolfe, D. E.; Dubé, W. P.; Ö ztürk, F.; Brock, C. A.; Grossberg, N.; Lefer, B.; Lerner, B.; Middlebrook, A. M.; Roberts, J. M. Understanding the Role of the Ground Surface in HONO Vertical Structure: High Resolution Vertical Profiles During NACHTT-11. J. Geophys. Res. Atmos. 2013, 118, 1−17. (9) Oswald, R.; Ermel, M.; Hens, K.; Novelli, A.; Ouwersloot, H. G.; Paasonen, P.; Petäjä, T.; Sipilä, M.; Keronen, P.; Bäck, J.; Königstedt, R.; Hosaynali Beygi, Z.; Fischer, H.; Bohn, B.; Kubistin, D.; Harder, H.; Martinez, M.; Williams, J.; Hoffmann, T.; Trebs, I.; Sörgel, M. A comparison of HONO budgets for two measurement heights at a field station within the boreal forest in Finland. Atmos. Chem. Phys. 2015, 15, 799−813. (10) Zhou, X.; Beine, H. J.; Honrath, R. E.; Fuentes, J. D.; Simpson, W.; Shepson, P. B.; Bottenheim, J. W. Snowpack photochemical production of HONO: a major source of OH in the Arctic boundary layer in springtime. Geophys. Res. Lett. 2001, 28, 4087−4090. (11) Finlayson-Pitts, B. J.; Wingen, L. M.; Sumner, A. L.; Syomin, D.; Ramazan, K. A. The heterogeneous hydrolysis of NO2 in laboratory systems and in outdoor and indoor atmospheres: an integrated mechanism. Phys. Chem. Chem. Phys. 2003, 5, 223−242. (12) Zhou, X.; Gao, H.; He, Y.; Huang, G.; Bertman, S. B.; Civerolo, K.; Schwab, J. Nitric acid photolysis on surfaces in low-NOx environments: significant atmospheric implications. Geophys. Res. Lett. 2003, 30, 2217. (13) Stemmler, K.; Ammann, M.; Donders, C.; Kleffmann, J.; George, C. Photosensitized reduction of nitrogen dioxide on humic acid as a source of nitrous acid. Nature 2006, 440, 195−198. (14) Kleffmann, J. Daytime sources of nitrous acid (HONO) in the atmospheric boundary layer. ChemPhysChem 2007, 8, 1137−1144. (15) Amoroso, A.; Domine, F.; Esposito, G.; Morin, S.; Savarino, J.; Nardino, M.; Montagnoli, M.; Bonneville, J. M.; Clement, J. C.; Ianniello, A.; Beine, H. J. Microorganisms in Dry Polar Snow Are Involved in the Exchanges of Reactive Nitrogen Species with the Atmosphere. Environ. Sci. Technol. 2010, 44, 714−719. (16) Oswald, R.; Behrendt, T.; Ermel, M.; Wu, D.; Su, H.; Cheng, Y.; Breuninger, C.; Moravek, A.; Mougin, E.; Delon, C.; Loubet, B.; Pommerening-Röser, A.; Sörgel, M.; Pöschl, U.; Hoffmann, T.; Andreae, M. O.; Meixner, F. X.; Trebs, I. HONO Emissions from Soil Bacteria as a Major Source of Atmospheric Reactive Nitrogen. Science 2013, 341, 1233−1235. (17) Su, H.; Cheng, Y.; Oswald, R.; Behrendt, T.; Trebs, I.; Meixner, F. X.; Andreae, M. O.; Cheng, P.; Zhang, Y.; Pöschl, U. Soil Nitrite as a Source of Atmospheric HONO and OH Radicals. Science 2011, 333, 1616−1618. (18) Stieglmeier, M.; Alves, R. E.; Schleper, C., The phylum Thaumarchaeota. In The Prokaryotes; Rosenberg, E., DeLong, E., Lory, S., Stackebrandt, E., Thompson, F., Eds.; Springer: Berlin Heidelberg, 2014; pp 347−362. (19) Koops, H.-P.; Purkhold, U.; Pommerening-Rö s er, A.; Timmermann, G.; Wagner, M., The lithoautotrophic ammoniaoxidizing bacteria. In The Prokaryotes; Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.-H., Stackebrandt, E., Eds.; Springer: New York, 2006; pp 778−811. (20) Abeliovich, A. The nitrite oxidizing bacteria. In The Prokaryotes, Dworkin, M., Falkow, S., Rosenberg, E., Schleifer, K.-H., Stackebrandt, E., Eds.; Springer: New York, 2006; pp 861−872. (21) Leininger, S.; Urich, T.; Schloter, M.; Schwark, L.; Qi, J.; Nicol, G. W.; Prosser, J. I.; Schuster, S. C.; Schleper, C. Archaea predominate among ammonia-oxidizing prokaryotes in soils. Nature 2006, 442, 806−809. (22) Bates, S. T.; Berg-Lyons, D.; Caporaso, J. G.; Walters, W. A.; Knight, R.; Fierer, N. Examining the global distribution of dominant archaeal populations in soil. ISME J. 2011, 5, 908−917.

ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.est.5b00838. Additional descriptions of the CEAS system; gene sequencing, bioinformatics and data reduction; sample descriptions and soil properties; a list of OTUs and their description for the most abundant microbes, AOA, AOB, and NOB (PDF)

■

AUTHOR INFORMATION

Corresponding Author Environ. Sci. Technol. Downloaded from pubs.acs.org by UNIV OF NEBRASKA-LINCOLN on 08/28/15. For personal use only.

*Phone: +1 (812) 855-6525; e-mail: JDRaff@indiana.edu. Present Address ∇

Department of Chemistry, Smith College, Northampton, Massachusetts 01063

Author Contributions ⊥

N.K.S. and U.M.E.S. contributed equally.

Notes

The authors declare no competing financial interest.

■

ACKNOWLEDGMENTS It is an honor to dedicate this paper to Professor Ronald Hitesmentor, colleague, coauthor, and friendin celebration of his immense contributions to the field of environmental chemistry. His pioneering applications of mass spectrometry to measuring trace organic chemicals in environmental matrices has yielded unparalleled advances in our understanding of the chemical fate and transport of pollutants in the environment. We look forward to many more years of collaboration and his inspiring contributions to the field. This work was funded by Indiana University (Office of the Vice Provost for Research, the School of Public and Environmental Affairs), the National Science Foundation (CAREER Award, AGS-1352375 to J.D.R.), an EPA STAR Graduate Fellowship (to N.K.S.), and an NSF Graduate Fellowship (to M.A.D.). We are grateful to Kathryn Fledderman and Laura Moore-Shay for assistance with the genomic work.

■

REFERENCES

(1) Finlayson-Pitts, B. J.; Pitts, J. N., Jr. Chemistry of the Upper and Lower AtmosphereTheory, Experiments, and Applications; Academic Press: San Diego, 2000; p 969. (2) Perner, D.; Platt, U. Detection of nitrous acid in the atmosphere by differential optical absorption. Geophys. Res. Lett. 1979, 6, 917−920. (3) Platt, U.; Perner, D.; Harris, G. W.; Winer, A. M.; Pitts, J. N., Jr. Observations of nitrous acid in an urban atmosphere by differential optical absorption. Nature 1980, 285, 312−314. (4) Kleffmann, J.; Kurtenbach, R.; Lörzer, J.; Wiesen, P.; Kalthoff, N.; Vogel, B.; Vogel, H. Measured and simulated vertical profiles of nitrous acidPart I: Field measurements. Atmos. Environ. 2003, 37, 2949−2955. (5) Wong, K. W.; Oh, H.-J.; Lefer, B. L.; Rappenglück, B.; Stutz, J. Vertical profiles of nitrous acid formation in the nocturnal urban atmosphere of Houston, TX. Atmos. Chem. Phys. 2011, 11, 3595− 3609. (6) Sörgel, M.; Trebs, I.; Serafimovich, A.; Moravek, A.; Held, A.; Zetzsch, C. Simultaneous HONO measurements in and above a forest canopy: influence of turbulent exchange on mixing ratio differences. Atmos. Chem. Phys. 2011, 11, 841−855. (7) Wong, K. W.; Tsai, C.; Lefer, B.; Haman, C.; Grossberg, N.; Brune, W. H.; Ren, X.; Luke, W.; Stutz, J. Daytime HONO vertical H


Article


Environmental Science & Technology (23) Taylor, A. E.; Zeglin, L. H.; Dooley, S.; Myrold, D. D.; Bottomley, P. J. Evidence for different contributions of archaea and bacteria to the ammonia-oxidizing potential of diverse Oregon soils. Appl. Environ. Microbiol. 2010, 76, 7691−8. (24) Martens-Habbena, W.; Berube, P. M.; Urakawa, H.; de la Torre, J. R.; Stahl, D. A. Ammonia oxidation kinetics determine niche separation of nitrifying Archaea and Bacteria. Nature 2009, 461, 976− 979. (25) Alves, R. J. E.; Wanek, W.; Zappe, A.; Richter, A.; Svenning, M. M.; Schleper, C.; Urich, T. Nitrification rates in Arctic soils are associated with functionally distinct populations of ammonia-oxidizing archaea. ISME J. 2013, 7, 1620−1631. (26) Stieglmeier, M.; Mooshammer, M.; Kitzler, B.; Wanek, W.; Zechmeister-Boltenstern, S.; Richter, A.; Schleper, C. Aerobic nitrous oxide production through N-nitrosating hybrid formation in ammoniaoxidizing archaea. ISME J. 2014, 8, 1135−46. (27) Zhang, L.-M.; Hu, H.-W.; Shen, J.-P.; He, J.-Z. Ammoniaoxidizing archaea have more important role than ammonia-oxidizing bacteria in ammonia oxidation of strongly acidic soils. ISME J. 2012, 6, 1032−1045. (28) Davidson, E. A. Sources of Nitric Oxide and Nitrous Oxide following Wetting of Dry Soil. Soil Sci. Soc. Am. J. 1992, 56, 95−102. (29) van Dijk, S. M.; Gut, A.; Kirkman, G. A.; Gomes, B. M.; Meixner, F. X.; Andreae, M. O. Biogenic NO emissions from forest and pasture soils: Relating laboratory studies to field measurements. J. Geophys. Res. 2002, 107, 8058. (30) Firestone, M. K.; Davidson, E. A. Microbiological basis of NO and N2O Production and Consumption in Soil; John Wiley & Sons Ltd.: Chichester, 1989; p 7−21. (31) Stutz, J.; Alicke, B.; Ackermann, R.; Geyer, A.; Wang, S. H.; White, A. B.; Williams, E. J.; Spicer, C. W.; Fast, J. D. Relative humidity dependence of HONO chemistry in urban areas. J. Geophys. Res. 2004, 109, D03307. (32) Riordan, E.; Minogue, N.; Healy, D.; O’Driscol, P.; Sodeau, J. R. Spectroscopic and Optimization Modeling Study of Nitrous Acid in Aqueous Solution. J. Phys. Chem. A 2005, 109, 779−786. (33) Donaldson, M. C.; Berke, A. E.; Raff, J. D. Uptake of Gas Phase Nitrous Acid onto Boundary Layer Soil Surfaces. Environ. Sci. Technol. 2013, 48, 11991−12001. (34) VandenBoer, T. C.; Markovic, M. Z.; Sanders, J. E.; Ren, X.; Pusede, S. E.; Browne, E. C.; Cohen, R. C.; Zhang, L.; Thomas, J.; Brune, W. H.; Murphy, J. G. Evidence for a nitrous acid (HONO) reservoir at the ground surface in Bakersfield, CA, during CalNex 2010. J. Geophys. Res.-Atmos. 2014, 119, 2013JD020971. (35) VandenBoer, T. C.; Young, C. J.; Talukdar, R. K.; Markovic, M. Z.; Brown, S. S.; Roberts, J. M.; Murphy, J. G. Nocturnal loss and daytime source of nitrous acid through reactive uptake and displacement. Nat. Geosci. 2014, 8, 55−60. (36) Donaldson, M. A.; Bish, D. L.; Raff, J. D. Soil surface acidity plays a determining role in the atmospheric-terrestrial exchange of nitrous acid. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 18472−18477. (37) Kubota, M.; Asami, T. Source of Nitrous Acid Volatilized from Upland Soils. Soil Sci. Plant Nutr. 1985, 31, 35−42. (38) Maljanen, M.; Yli-Pirilä, P.; Hytönen, J.; Joutsensaari, J.; Martikainen, P. J. Acidic northern soils as sources of atmospheric nitrous acid (HONO). Soil Biol. Biochem. 2013, 67, 94−97. (39) Wu, D.; Kampf, C. J.; Pöschl, U.; Oswald, R.; Cui, J.; Ermel, M.; Hu, C.; Trebs, I.; Sörgel, M. Novel Tracer Method To Measure Isotopic Labeled Gas-Phase Nitrous Acid (HO15NO) in Biogeochemical Studies. Environ. Sci. Technol. 2014, 48, 8021−8027. (40) Hendershot, W. H.; Lalande, H.; Duquette, M., Soil Reaction and Exchangeable Acidity. In Soil Sampling and Methods of Analysis, 2nd ed.; Carter, M. R.; Gregorich, E. G., Eds.; Canadian Society of Soil Science: 2008; pp 173−178. (41) Keeney, D. R.; Nelson, D. W., Nitrogen-Inorganic Forms. Method 33-3.2. In Methods of Soil Analysis. Part 2. Chemical and Microbiological Properties, 2nd ed.; Page, A. L., Ed. ASA, SSA: Madison, WI, 1982; pp 643−698.

(42) Washenfelder, R. A.; Langford, A. O.; Fuchs, H.; Brown, S. S. Measurement of glyoxal using an incoherent broadband cavity enhanced absorption spectrometer. Atmos. Chem. Phys. 2008, 8, 7779−7793. (43) Scharko, N. K.; Berke, A. E.; Raff, J. D. Release of Nitrous Acid and Nitrogen Dioxide from Nitrate Photolysis in Acidic Aqueous Solutions. Environ. Sci. Technol. 2014, 48, 11991−12001. (44) Kebede, M. A.; Scharko, N. K.; Appelt, L. E.; Raff, J. D. Formation of nitrous acid during ammonia photooxidation on TiO2 under atmospherically relevant conditions. J. Phys. Chem. Lett. 2013, 4, 2618−2623. (45) Gilbert, J. A.; Jansson, J. K.; Knight, R. The Earth Microbiome project: successes and aspirations. BMC Biol. 2014, 12, 69. (46) The Earth Microbiome Project: 16S rRNA Amplification Protocol. http://www.earthmicrobiome.org/emp-standard-protocols/ 16s/ (accessed February 11, 2015). (47) Quast, C.; Pruesse, E.; Yilmaz, P.; Gerken, J.; Schweer, T.; Yarza, P.; Peplies, J.; Glockner, F. O. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013, 41, D590−6. (48) Caporaso, J. G.; Lauber, C. L.; Walters, W. A.; Berg-Lyons, D.; Lozupone, C. A.; Turnbaugh, P. J.; Fierer, N.; Knight, R. Global patterns of 16S rRNA diversity at a depth of millions of sequences per sample. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 4516−4522. (49) Belser, L.; Schmidt, E. Inhibitory effect of nitrapyrin on three genera of ammonia-oxidizing nitrifiers. Appl. Environ. Microbiol. 1981, 41, 819−821. (50) Bremner, J.; Breitenbeck, G.; Blackmer, A. Effect of nitrapyrin on emission of nitrous oxide from soil fertilized with anhydrous ammonia. Geophys. Res. Lett. 1981, 8, 353−356. (51) Powell, S. J.; Prosser, J. I. Inhibition of ammonium oxidation by nitrapyrin in soil and liquid culture. Appl. Environ. Microbiol. 1986, 52, 782−787. (52) Angelini, M. M.; Garrard, R. J.; Rosen, S. J.; Hinrichs, R. Z. Heterogeneous reactions of gaseous HNO3 and NO2 on the clay minerals kaolinite and pyrophyllite. J. Phys. Chem. A 2007, 111, 3326− 3335. (53) Usher, C. R.; Michel, A. E.; Grassian, V. H. Reactions on mineral dust. Chem. Rev. 2003, 103, 4883−4940. (54) Pester, M.; Rattei, T.; Flechl, S.; Grongroft, A.; Richter, A.; Overmann, J.; Reinhold-Hurek, B.; Loy, A.; Wagner, M. amoA-based consensus phylogeny of ammonia-oxidizing archaea and deep sequencing of amoA genes from soils of four different geographic regions. Environ. Microbiol. 2012, 14, 525−539. (55) Pester, M.; Schleper, C.; Wagner, M. The Thaumarchaeota: an emerging view of their phylogeny and ecophysiology. Curr. Opin. Microbiol. 2011, 14, 300−306. (56) Stein, L. Y.; Klotz, M. G. Nitrifying and denitrifying pathways of methanotrophic bacteria. Biochem. Soc. Trans. 2011, 39, 1826−1831. (57) Lücker, S.; Daims, H. The Prokaryotes: Deltaproteobacteria and Epsilonproteobacteria; Springer: Berlin Heidelberg, 2014. (58) Ke, X.; Angel, R.; Lu, Y.; Conrad, R. Niche differentiation of ammonia oxidizers and nitrite oxidizers in rice paddy soil. Environ. Microbiol. 2013, 15, 2275−92. (59) Cebron, A.; Garnier, J. Nitrobacter and Nitrospira genera as representatives of nitrite-oxidizing bacteria: detection, quantification and growth along the lower Seine River (France). Water Res. 2005, 39, 4979−4992. (60) Paavolainen, L.; Kitunen, V.; Smolander, A. Inhibition of nitrification in forest soil by monoterpenes. Plant Soil 1998, 205, 147− 154. (61) Ward, B. B.; Courtney, K. J.; Langenheim, J. H. Inhibition of Nitrosomonas europaea by monoterpenes from coastal redwood (Sequoia sempervirens) in whole-cell studies. J. Chem. Ecol. 1997, 23, 2583−2598. (62) White, C. S. Monoterpenes-Their Effects on Ecosystem Nutrient Cycling. J. Chem. Ecol. 1994, 20, 1381−1406. I


Article



bacterial community structure at the continental scale. Appl. Environ. Microbiol. 2009, 75, 5111−20. (84) Fierer, N.; Leff, J.; Adams, B.; Nielsen, U.; Bates, S.; Lauber, C.; Owens, S.; Gilbert, J.; Wall, D.; Caporaso, J. Cross-biome metagenomic analyses of soil microbial communities and their functional attributes. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 21390−21395. (85) Nicol, G. W.; Leininger, S.; Schleper, C.; Prosser, J. I. The influence of soil pH on the diversity, abundance and transcriptional activity of ammonia oxidizing archaea and bacteria. Environ. Microbiol. 2008, 10, 2966−78. (86) Lehtovirta-Morley, L. E.; Stoecker, K.; Vilcinskas, A.; Prosser, J. I.; Nicol, G. W. Cultivation of an obligate acidophilic ammonia oxidizer from a nitrifying acid soil. Proc. Natl. Acad. Sci. U. S. A. 2011, 108, 15892−15897. (87) Hatzenpichler, R.; Lebedeva, E. V.; Spieck, E.; Stoecker, K.; Richter, A.; Daims, H.; Wagner, M. A moderately thermophilic ammonia-oxidizing crenarchaeote from a hot spring. Proc. Natl. Acad. Sci. U. S. A. 2008, 105, 2134−2139. (88) Harrison, R. M.; Kitto, A.-M. N. Evidence for a Surface Source of Atmospheric Nitrous Acid. Atmos. Environ. 1994, 28, 1089−1094. (89) Stutz, J.; Alicke, B.; Neftel, A., Nitrous acid formation in the urban atmosphere: Gradient measurements of NO2 and HONO over grass in Milan, Italy. J. Geophys. Res. 2002, 107, DOI:10.1029/ 2001JD000390. (90) Zhou, X.; Zhang, N.; TerAvest, M.; Tang, D.; Hou, J.; Bertman, S.; Alaghmand, M.; Shepson, P. B.; Carroll, M. A.; Griffith, S.; Dusanter, S.; Stevens, P. S. Nitric acid photolysis on forest canopy surface as a source for tropospheric nitrous acid. Nat. Geosci. 2011, 4, 440−443. (91) Ren, X.; Sanders, J. E.; Rajendran, A.; Weber, R. J.; Goldstein, A. H.; Pusede, S. E.; Browne, E. C.; Min, K. E.; Cohen, R. C. A relaxed eddy accumulation system for measuring vertical fluxes of nitrous acid. Atmos. Meas. Tech. 2011, 4, 2093−2103. (92) Raciti, S. M.; Groffman, P. M.; Fahey, T. J. Nitrogen retention in urban lawns and forests. Ecol Appl. 2008, 18, 1615−26. (93) Raciti, S. M.; Burgin, A. J.; Groffman, P. M.; Lewis, D. N.; Fahey, T. J. Denitrification in suburban lawn soils. J. Environ. Qual. 2011, 40, 1932−40. (94) Groffman, P. M.; Boulware, N. J.; Zipperer, W. C.; Pouyat, R. V.; Band, L. E.; Colosimo, M. F. Soil nitrogen cycle processes in urban riparian zones. Environ. Sci. Technol. 2002, 36, 4547−4552. (95) Lorenz, K.; Kandeler, E. Microbial biomass activities in urban soils in two consecutive years. J. Plant Nutr. Soil Sci. 2006, 169, 799− 808. (96) Raciti, S. M.; Groffman, P. M.; Jenkins, J. C.; Pouyat, R. V.; Fahey, T. J.; Pickett, S. T. A.; Cadenasso, M. L. Nitrate production and availability in residential soils. Ecol. Appl. 2011, 21, 2357−2366. (97) Bettez, N.; Marino, R.; Howarth, R.; Davidson, E. Roads as nitrogen deposition hot spots. Biogeochemistry 2013, 114, 149−163. (98) Czader, B. H.; Choi, Y.; Li, X.; Alvarez, S.; Lefer, B. Impact of updated traffic emissions on HONO mixing ratios simulated for urban site in Houston, Texas. Atmos. Chem. Phys. 2015, 15, 1253−1263.

(63) Austin, A.; Yahdjian, L.; Stark, J.; Belnap, J.; Porporato, A.; Norton, U.; Ravetta, D.; Schaeffer, S. Water pulses and biogeochemical cycles in arid and semiarid ecosystems. Oecologia 2004, 141, 221−235. (64) Kim, D. G.; Vargas, R.; Bond-Lamberty, B.; Turetsky, M. R. Effects of soil rewetting and thawing on soil gas fluxes: a review of current literature and suggestions for future research. Biogeosciences 2012, 9, 2459−2483. (65) Placella, S. A.; Brodie, E. L.; Firestone, M. K. Rainfall-induced carbon dioxide pulses result from sequential resuscitation of phylogenetically clustered microbial groups. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 10931−10936. (66) Placella, S. A.; Firestone, M. K. Transcriptional response of nitrifying communities to wetting of dry soil. Appl. Environ. Microbiol. 2013, 79, 3294−3302. (67) Gelfand, I.; Yakir, D. Influence of nitrite accumulation in association with seasonal patterns and mineralization of soil nitrogen in a semi-arid pine forest. Soil Biol. Biochem. 2008, 40, 415−424. (68) Hudman, R. C.; Moore, N. E.; Mebust, A. K.; Martin, R. V.; Russell, A. R.; Valin, L. C.; Cohen, R. C. Steps towards a mechanistic model of global soil nitric oxide emissions: implementation and space based-constraints. Atmos. Chem. Phys. 2012, 12, 7779−7795. (69) Davidson, E. A. Pulses of Nitric Oxide and Nitrous Oxide Flux following Wetting of Dry Soil: An Assessment of Probable Sources and Importance Relative to Annual Fluxes. Ecol. Bull. 1992, 149−155. (70) Dividson, E. A.; Stark, J. M.; Firestone, M. K. Microbial Production and Consumption of Nitrate in an Annual Grassland. Ecology 1990, 71, 1968−1975. (71) Flowers, T. H.; O’Callaghan, J. R. Nitrification in soils incubated with pig slurry or ammonium sulphate. Soil Biol. Biochem. 1983, 15, 337−342. (72) Hopkins, D. W.; O’Donnell, A. G.; Shiel, R. S. The effect of fertilisation on soil nitrifier activity in experimental grassland plots. Biol. Fertil. Soils 1988, 5, 344−349. (73) De Boer, W.; Gunnewiek, P.; Veenhuis, M.; Bock, E.; Laanbroek, H. Nitrification at Low pH by Aggregated Chemolithotrophic Bacteria. Appl. Environ. Microbiol. 1991, 57, 3600−3604. (74) De Boer, W.; Kowalchuk, G. Nitrification in acid soils: microorganisms and mechanisms. Soil Biol. Biochem. 2001, 33, 853−866. (75) Hatzenpichler, R. Diversity, physiology, and niche differentiation of ammonia-oxidizing archaea. Appl. Environ. Microbiol. 2012, 78, 7501−7510. (76) Stevens, R.; Laughlin, R.; Malone, J. Soil pH affects the processes reducing nitrate to nitrous oxide and di-nitrogen. Soil Biol. Biochem. 1998, 30, 1119−1126. (77) Ste-Marie, C.; Pare, D. Soil, pH and N availability effects on net nitrification in the forest floors of a range of boreal forest stands. Soil Biol. Biochem. 1999, 31, 1579−1589. (78) Stark, J. M.; Hart, S. C., High rates of nitrification and nitrate turnover in undisturbed coniferous forests. Nature 1997385, 61− 64.10.1038/385061a0 (79) Van Cleemput, O.; Baert, L. Nitrite stability influenced by iron compounds. Soil Biol. Biochem. 1983, 15, 137−140. (80) Coby, A. J.; Picardal, F. W. Inhibition of NO3− and NO2− reduction by microbial Fe(III) reduction: evidence of a reaction between NO2− and cell surface-bound Fe2+. Appl. Environ. Microbiol. 2005, 71, 5267−5274. (81) Guerbois, D.; Ona-Nguema, G.; Morin, G.; Abdelmoula, M.; Laverman, A. M.; Mouchel, J.-M.; Barthelemy, K.; Maillot, F.; Brest, J. Nitrite Reduction by Biogenic Hydroxycarbonate Green Rusts: Evidence for Hydroxy-nitrite Green Rust Formation as an Intermediate Reaction Product. Environ. Sci. Technol. 2014, 48, 4505−4514. (82) Sörgel, M.; Trebs, I.; Wu, D.; Held, A. A comparison of measured HONO uptake and release with calculated source strengths in a heterogeneous forest environment. Atmos. Chem. Phys. Discuss. 2015, 15, 2119−2155. (83) Lauber, C. L.; Hamady, M.; Knight, R.; Fierer, N. Pyrosequencing-based assessment of soil pH as a predictor of soil J


Combined Flux Chamber and Genomics Approach ... - ACS Publications

Recommend Documents