Evidence for Different Contributions of Archaea and Bacteria to the Ammonia-Oxidizing Potential of Diverse Oregon Soils

Chemicals.ATU, N-tris(hydroxymethyl)methyl-2-aminoethanesulfonic acid (TES) buffer, neomycin trisulfate salt, nystatin, dimethyl sulfoxide (DMSO) (99.5%), actinomycin D, and NH₄Cl were obtained from Sigma (St. Louis, MO). Acetylene was obtained from Airgas (Radnor, PA); cycloheximide (96.4%), kanamycin sulfate, and gentamicin sulfate were obtained from EMD Biosciences, Inc. (La Jolla, CA); and emetine was purchased from TCI America (Portland, OR). The fungicide Abound, with the active ingredient methyl-(E)-2-{2-[6-(cyanophenoxy)pyrimidin-4-yloxy]phenyl}-3-methoxyacrylate (azoxystrobin), was obtained from Syngenta Crop Protection, Inc. (Greensboro, NC), and Szechrome NAS reagent was obtained from Polysciences, Inc. (Warrington, PA).

Soils.After the removal of litter or sod, mineral soil samples (0 to 10 cm) were collected from sites along a west-to-east transect of different land uses approximately 6 miles (9.5 km) in length and located 10 miles (16 km) north of Corvallis, OR (44.7°N, 123.3°W). Soils were taken from three sites under mature red alder stands (forest) in the uplands (∼350-m elevation) of the MacDonald-Dunn Forest (Jory-Gelderman series, Paleo/Haplohumults [http://soils.usda.gov/technical/classification/tax_keys/], pH 6.0 to 6.3) from three sites under permanent grass pastures situated on a toe slope below the forest (∼125-m elevation) on the Oregon State University (OSU) Beef Farm (Dixonville-Gellatly-Witham complex, Argi/Haploxerolls, pH 5.7 to 6.4) and from three cropped fields (under wheat or barley at the time of sampling) and three fallowed fields (unfertilized and uncultivated since 1990 and colonized by volunteer grasses and forbs, which are mowed twice yearly) at the OSU Hyslop Field Research Laboratory (Woodburn series, Argixerolls, pH 5.9 to 6.5) on the Willamette Valley floor (∼85-m elevation). Four to five soil samples were recovered from each of the three locations that were spaced between 100 m and ∼1,000 m apart in each of the forest, pasture, cropped, and fallowed areas. The samples at each location were composited, thoroughly mixed in the field, and brought to the laboratory, where they were sieved (4.75 mm) and stored at 4°C. Samples of soil (5 to 10 g) were oven dried to determine the water content. The levels of net readily mineralizable nitrogen (N Min) were determined for soils adjusted to ∼60% of the water-holding capacity, incubated at 30°C, and sampled at 7, 14, 21, and 30 days, and the accumulations of ammonium (NH₄⁺) plus nitrate (NO₃⁻) were measured. Physical and chemical properties of the soil samples were determined by standard procedures of the Central Analytical Services Laboratory, Department of Crop and Soil Science, Oregon State University.

NP.Nitrification potential (NP) assays are used primarily to characterize the potential activity of the nitrifying population under conditions of nonlimiting NH₃ and O₂ and neutral pH (51). Soils were removed from 4°C storage and preincubated field moist at room temperature (22°C ± 2°C) for 2 days prior to experimental use. The fallowed and cropped soils from Hyslop Farm had a low water content (≤11%, wt/wt); therefore, deionized water was added (1 ml/15 g soil) before preincubation to achieve a water content that approximates field capacity (18%, wt/wt). For NP assays, 5-g portions of moist soil were added to 50-ml aliquots of 30 mM TES, buffered to pH 7.2 with 10 M KOH, in 150-ml bottles with black phenolic caps fitted with gray butyl stoppers. TES buffer (30 mM) was used because its pK_a is pH 7.4, it did not extract humic compounds from the soil, and its buffering capacity was sufficient to maintain the soil slurries at pH 6.8 to 7.0 for the duration of the assays. Preliminary experiments showed that 1 mM NH₄⁺ was a saturating, noninhibiting concentration, and all NP assays included 1 mM supplemental NH₄⁺ (as NH₄Cl) unless otherwise stated. Soil slurries were shaken at 200 rpm either in a Brunswick orbital constant-temperature shaker maintained at either 30°C or 40°C or on a bench-top orbital shaker at room temperature (22°C ± 2°C). At time intervals, samples (1.8 ml) of soil slurry were removed into microcentrifuge tubes and centrifuged for 3 min at 13.4 × 10³ × g, and nitrite (NO₂⁻) and NO₃⁻ concentrations in the supernatants were determined immediately or stored at −20°C until the samples could be processed. NO₂⁻ and NO₃⁻ concentrations were calculated using the oven-dried weight of the soil used in the assay. Background NO₂⁻ and NO₃⁻ concentrations were measured after soil slurries were shaken for 15 min. NP rates were determined as NO₂⁻ plus NO₃⁻ accumulations over 24 h. Although background NO₂⁻ was undetected in any of the soils, NO₂⁻ accumulated to various fractions of the total NO₂⁻ plus NO₃⁻ in actively nitrifying soil slurries. NO₂⁻ concentrations were determined as described elsewhere (22). NO₃⁻ concentrations were determined by using either Szechrome reagent according to the manufacturer's instructions or an Astoria Pacific (Clackamas, OR) flow solution autoanalyzer. It was determined that both ATU and cycloheximide interfered with the Szechrome assay, and in those cases, the NO₃⁻ concentration was always determined by use of an autoanalyzer. In a number of experiments the NH₄⁺ concentration was also determined by using the flow solution autoanalyzer. In some cases, high concentrations of background soil NO₃⁻ were removed prior to NP assays as follows: 5-g portions of moist soils were dispensed into 40-ml centrifuge tubes with ∼30 ml of distilled water (dH₂O), vortexed briefly, and centrifuged at 7.5 ×10³ × g for 15 min, and the supernatant was carefully decanted. Soil pellets were resuspended in NP assay buffer and transferred into assay bottles. Preliminary experiments showed that pretreating the soils in this manner led to a minimal loss of NP.

The effects of bacterial and eukaryotic protein synthesis inhibitors, ATU, and fungicides on nitrification were determined by comparing nitrifications in the presence and absence of the individual compounds. During preliminary experiments with the relatively water-insoluble antibiotic actinomycin D, we found that nitrification in soil slurries was inhibited by extremely small volumes of DMSO and other solvents such as methanol, thereby preventing the use of actinomycin D concentrations of >200 μg/ml. Due to their insolubility in aqueous solutions, we did not pursue the use of antibiotics such as rifampin or chloramphenicol or fungicides such as triazoles. The water-soluble bacterial protein synthesis inhibitors neomycin, kanamycin, and gentamicin were utilized. Kanamycin and gentamicin have been proven effective in preventing the growth of the AOB Nitrosomonas europaea (13, 21, 56). The effects of the water-soluble eukaryotic protein synthesis inhibitors emetine and cycloheximide were evaluated. A stock of the nitrification inhibitor ATU was mixed immediately before use. The water-soluble fungicide nystatin (1.2 ml/50 ml; 12,000 units) or azoxystrobin (20 μl/50 ml; 13 mg of active ingredient) was used as recommended by the manufacturers.

RNP.For assays of the recovery of nitrification potential (RNP), 5-g portions of moist soil were dispensed into 50-ml aliquots of 30 mM TES buffer (pH 7.2) in 150-ml bottles with black phenolic caps fitted with gray butyl stoppers. Acetylene was added (0.025%, vol/vol, or 0.025 kPa) to the headspace for 6 h. From preliminary experiments it was determined that a 6-h acetylene exposure was sufficient to inactivate all NH₃ oxidation in the soils used in the study. Acetylene inhibition and RNP steps were carried out at 30°C with 1 mM supplemental NH₄⁺, unless otherwise indicated. These conditions in the absence of inhibitors were considered to be the standard RNP, designated RNP°. NP controls comprised soil suspensions to which acetylene was not added. An acetylene-containing control was also included to evaluate the possibility of heterotrophic nitrification. The removal of acetylene was achieved by placing the soil slurries under a vacuum (−100 kPa) and degassing. Analysis of headspace samples by gas chromatography with flame ionization detection (FID), as described previously (59), determined that degassing for 5 min under a vacuum was adequate to remove all traces of acetylene from soil slurries.

After degassing, all bottles were incubated with caps loosened to permit aeration. Although RNP varied with soil samples from different land use regimens, replicate soil samples from the same land use regimen responded similarly. In the case of pasture and cropped soil samples, the length of lag time was evaluated by determining the accumulation of NO₂⁻ plus NO₃⁻ at 6-h intervals for 24 to 30 h, followed by sampling at less frequent intervals for a total of 48 h. Fallowed soil had a lower NP, showed a longer lag time (∼24 to 30 h), and recovered more slowly. The RNPs were compared with the initial NP controls by assessing the amounts of NO₂⁻ plus NO₃⁻ that accumulated in the 24-h postlag period relative to the accumulation occurring between 0 and 24 h in the NP. From preliminary experiments we established there were no detrimental effects of degassing the NP controls after 6 h of incubation on the subsequent rate. Aqueous solutions of antibiotics, nitrification inhibitors, or fungicides were added after acetylene was evacuated. Preliminary experiments were carried out with a range of concentrations of the antibiotics to determine their minimum effective concentrations.

Extraction of nucleic acids and Q-PCR.DNA was extracted from 0.25 to 0.3 g dry soil by using a MoBio (Carlsbad, CA) PowerSoil extraction kit, and DNA was quantified by using a NanoDrop ND-1000 UV-Vis spectrophotometer (ThermoScientific, Rockwood, TN). Quantitative PCR (Q-PCR) of the archaeal and bacterial amoA genes was performed by using Brilliant II SYBR green master mix (Stratagene, La Jolla, CA) and an ABI 7500 real-time PCR system. Each 25-μl reaction mixture volume included 10 ng template DNA. Primers and thermal cycler protocols for both bacterial (primers amoA_1R and amoA_2F) and archaeal (primers Arch-amoAF and Arch-amoAR) amoA genes were described elsewhere previously (41, 48). Standard curves were constructed with 2.0 × 10¹ to 2.0 × 10⁻⁴ ng Nitrosomonas europaea genomic DNA (bacterial amoA [efficiency = 91 to 94%; R² = 0.996]) or 1.25 × 10¹ to 1.25 × 10⁻³ ng of a TOPO plasmid containing the “Candidatus Nitrosopumilus maritimus” strain SCM1 amoBAC gene insert (archaeal amoA [efficiency = 95 to 100%; R² = 0.994 to 0.995]). Each reaction was run in triplicate. Copy numbers were standardized by the mass of DNA and g dry soil analyzed for each sample and log₁₀ transformed before further analysis.

Statistics.We used one-way analysis of variance (ANOVA) with Bonferroni post hoc analysis using SPSS 11 for Mac to determine the significance of temperature effects within each incubation set. A single-tailed Student's t test assuming unequal variance was used to evaluate the effect of supplemental NH₄⁺ on NP and to determine the effect of inhibitors and temperature on RNP.

Soil properties.Due to their close geographic proximity, the soils taken along the landscape transect experience approximately the same precipitation and temperature regimen. Archived soil temperature data from the cropped and fallowed sites show annual maxima and minima of 37.2°C ± 1.5°C and 2.9°C ± 0.9°C, respectively (http://cropandsoil.oregonstate.edu/weather/archive). A suite of chemical and physical properties was determined for the soils (Table 1). While there were only small differences in pHs among the sites, other soil characteristics varied to a greater extent. Notably, high extractable P concentrations in cropped and fallowed soil samples reflect the legacy of P fertilization to promote crop yield, and the high levels of Mg, Na, Ca, and clay in the pasture soils are likely due to the landscape position (toe slope). The cropped soils had the least amount of total C; the adjacent fallowed soils had been out of production for 19 years and had C levels that were 1.9-fold greater than those of the cropped soils. The pasture soils had 2.1-fold-more C than the fallowed soils.

Levels of N that were readily mineralizable over 30 days (N Min) followed a trend similar to that of total C (Table 1). The fallowed soils had a 1.6-fold-greater N Min than the cropped soils, and the pasture soils had a 1.4-fold-greater N Min than the fallowed soils. The highest level of N Min was found in the forest soils and probably reflects the high N input by the N₂-fixing red alder trees at these sites. Q-PCR of amoA genes of AOA and AOB demonstrated the presence of both types of NH₃ oxidizers at all sites. Generally, the AOA and AOB amoA copy numbers/g soil were within the same order of magnitude, except for pasture soils, where the AOA amoA copy number/g soil was 2 orders of magnitude higher than the AOB amoA copy number (41.7 × 10⁶ and 0.4 × 10⁶ copies/g soil, respectively).

Properties of soil nitrification potentials (NPs).We conducted an initial screening of soil NPs that revealed that the nitrifier communities possessed different characteristics among the soils and thus provided distinct soil phenotypes for comparing the properties of the recovered NP.

(i) Effects of temperature.The NPs of the soils varied in their responses to temperature (Fig. 1). Pasture soils showed the greatest relative increase in NP at between 22°C and 30°C (3.5-fold), and NPs at 30°C and 40°C were significantly greater (P ≤ 0.01) than those at 22°C. Forest and cropped soils also showed increases in NPs between 22°C and 30°C, 1.8- and 1.4-fold, respectively, but showed significantly lower (P ≤ 0.01) levels of activity at 40°C than at 30°C. Cropped soils had only a small fraction (0.06) of their 30°C NP activity at 40°C, while forest soil showed approximately one-half of the 30°C activity at 40°C. At 22°C the NPs of fallowed soils were similar to those of pasture soils but did not increase significantly when incubated at 30°C. The response of fallowed soil NPs to 40°C was mixed: in two cases, the activity at 40°C was less than the NP at 30°C, whereas in one case, the activities were the same.

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FIG. 1.

Temperature responses of the nitrification potentials (NPs) of soil samples taken from three sites each of forest, pasture, cropped, and fallowed land. Mean values and standard deviations are shown (n = 3).

(ii) Effects of supplemental NH₄⁺.The NPs of all soils produced at 30°C were greater than the rates of net N mineralization over the first 7 days of the N Min study of the same soils (Fig. 2). When NPs were done without supplemental NH₄⁺, the accumulation of NO₂⁻ plus NO₃⁻ was significantly lower (P ≤ 0.01) in the forest, pasture, and cropped soils than in the presence of 1 mM supplemental NH₄⁺. However, fallowed soils produced the same NP with or without supplemental NH₄⁺ (P = 0.4). Whereas cropped soils demonstrated only 11% of the plus NH₄⁺ rate in the absence of supplemental NH₄⁺, forest and pasture soils supported NPs of 57 and 69%, respectively, of their rate with NH₄⁺. In all cases, during the 24-h time course, the concentrations of NH₄⁺ in NP assay solutions without supplemental NH₄⁺ were below the limit of accurate detection by the autoanalyzer (≤10 μM). Subsequently, NH₄⁺ began to accumulate as N mineralization responded to the incubation at 30°C.

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FIG. 2.

Comparison of whole-soil rates of net N mineralization and the NPs of soil slurries with and without supplemental NH₄⁺. The net N mineralization rate equals NH₄⁺ plus NO₃⁻ mineralized at 30°C over the first 7 days of the N Min study. Mean values and standard deviations are shown for three analytical replicates of a composite soil sample prepared from individual composite samples recovered from each of three field sites under the same land use conditions. ** indicates that values for NP plus 1 mM supplemental NH₄⁺ are significantly different from values for NP without added NH₄⁺ (P < 0.01).

(iii) Effect of ATU.The NPs of soils were measured in the presence of the nitrification inhibitor allyl thiourea (ATU). In the presence of 100 μM ATU, virtually all (92 to 95%) of the NPs of forest, cropped, and fallowed soils were inhibited, whereas pasture soils maintained 53% ± 6% of their NO₂⁻-plus-NO₃⁻-producing activity. At 40°C, pasture soils retained all of their NP activity at concentrations of ≤1 mM ATU (data not shown). The ATU insensitivity of pasture soil NPs was not due to soil concentrations of Cu²⁺ exceeding the chelating capacity of ATU, as the extractable Cu²⁺ level in 30 mM TES buffer was determined to be below the limits of analytical detection (<0.08 μM) (data not shown), and thus, the ATU level was far in excess of Cu²⁺ levels. Mg²⁺ and Ca²⁺ are divalent cations that could also potentially interfere with the Cu-chelating ability of ATU. The Ca²⁺ and Mg²⁺ concentrations in pasture soil slurry solutions were determined to be 0.73 and 0.48 mM, respectively. We obtained no evidence that the ATU sensitivity of cropped soil slurries could be reduced by raising their Ca²⁺ and Mg²⁺ concentrations to pasture soil slurry levels (data not shown).

Properties of acetylene inactivation and RNP.NPs of all soils were completely inhibited by exposure to 0.025% (vol/vol) headspace acetylene (8 μM aqueous) for 6 h (Fig. 3), indicating that the oxidation of NH₃ in the soils was mediated by a monooxygenase(s) and that heterotrophic nitrification was negligible. After acetylene was removed and slurries were further incubated at 30°C, a significant accumulation of NO₂⁻ plus NO₃⁻ could be detected in pasture, cropped, and fallowed soils 12, 18, and 24 to 30 h, respectively, after acetylene removal (Fig. 3). During the 24-h time interval immediately following the acetylene-induced lag, pasture, cropped, and fallowed soils regained 100% ± 39%, 57% ± 36%, and 82% ± 30%, respectively, of the NO₂⁻ plus NO₃⁻ produced by their respective NP controls. For unknown reasons, soil slurries from the three forest sites did not consistently demonstrate recovered nitrification potential (RNP) regardless of the incubation temperature (data not shown). The RNP of forest soils did not consistently recover even when time of exposure of slurries to acetylene was reduced to 1 h or when slurries were supplemented with either 5 mM NH₄⁺, trace elements (including Cu²⁺) from standard AOB growth medium (44), or 20 mg/liter of organic supplements, yeast extract, or tryptic soy broth.

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FIG. 3.

Comparison of the time courses of NPs and recovered nitrification potentials (RNPs) from representative examples of pasture, cropped, and fallowed soils at 30°C. Error bars indicate the standard deviations of data from three analytical replicates. RNPs were treated with 0.025% (vol/vol) acetylene at time zero. Arrows indicate the time at which acetylene was removed (6 h).

The time delay in the RNP of the soils indicated that de novo protein synthesis might be required to replace irreversibly inactivated AMO before the resumption of NO₂⁻ and NO₃⁻ production could occur. Indeed, the RNPs of cropped soils were 90% ± 10% and 80% ± 0% prevented by 800 μg/ml kanamycin and gentamicin, respectively (Table 2). The antibiotics did not have an effect on the non-acetylene-treated NP control of cropped soil for 18 h. After 18 h, the production of NO₂⁻ and NO₃⁻ ceased (data not shown), indicating that the antibiotics were not inhibitors of nitrification per se and were likely preventing de novo protein synthesis. In the fallowed soils, only fractions of RNP were prevented by either kanamycin or gentamicin (50% and 10%, respectively), but a combination of both antibiotics had a greater effect (60% ± 30%). In complete contrast, and over a wide range of concentrations (≤800 μg/ml), the bacterial protein synthesis inhibitors neomycin (data not shown) and kanamycin and gentamicin, singly or in combination, had virtually no effect on the RNP of pasture soils (Table 2). In addition, kanamycin had no effect on the NP of pasture soils over 0 to 48 h of incubation (data not shown). Similar results were obtained with samples of pasture soil taken at other times of the year (May and December) and from another series of three pasture sites located one-half mile further upslope (data not shown).

We examined the properties of the RNP from each soil sample to determine to what extent they matched those of the NPs (Table 3). Patterns of temperature response and ATU sensitivity were similar to those demonstrated for the NP controls. The RNPs of pasture and fallowed soils occurred at both 30°C and 40°C, while in cropped soils, RNP did not occur at 40°C. Whereas RNPs of both cropped and fallowed soils were prevented by 100 μM ATU, the RNP of pasture soil was inhibited by only ∼50%, and at 40°C, the RNP of pasture soil was completely insensitive to ATU concentrations of ≤1 mM (data not shown). Furthermore, cropped soil RNP did not occur in the absence of supplemental NH₄⁺, whereas RNPs of both pasture and fallowed soils occurred in the absence of supplemental NH₄⁺ (80% ± 20% and 40% ± 20% of their RNP controls, respectively) at soil solution concentrations of NH₄⁺ too low to be accurately measured by the autoanalyzer (≤10 μM). All of the RNPs were completely inactivated by the addition of acetylene (data not shown).

Because of the complete insensitivity of the RNP of pasture soil to bacterial protein synthesis inhibitors, we evaluated the effects of eukaryotic protein synthesis inhibitors and especially ones that have been reported to be effective on archaea. No inhibition of RNPs was observed for pasture soil slurries treated with the eukaryotic transcriptional inhibitor actinomycin D (200 μg/ml) (data not shown), and the protein synthesis inhibitor emetine (800 μg/ml) had no effect on pasture, fallow, or cropped soils (data not shown). Furthermore, neither of the fungicides nystatin and azoxystrobin had an effect on the RNP of pasture or fallowed soils (data not shown). Despite the ineffectiveness of the above-mentioned compounds, an unpublished observation from our laboratory has shown that the growth of the AOA “Candidatus Nitrosopumilus maritimus” strain SCM1 is strongly inhibited by 200 μg/ml of the eukaryotic protein synthesis inhibitor cycloheximide, while the growth of Nitrosomonas europaea is insensitive (N. Vajrala, personal communication). We obtained reproducible evidence for a negative effect of 600 to 800 μg/ml cycloheximide on the RNP of pasture soil (0.6 ± 0.2 of their RNP controls); however, cycloheximide had a similarly negative effect on the RNP of cropped and fallowed soils as well (0.5 ± 0.4 and 0.3 ± 0.1 of their RNP controls, respectively). A wide range of cycloheximide concentrations (25 to 600 μg/ml) was shown to have no immediate effect (0 to 12 h) on the NPs of either pasture or cropped soils, suggesting that its effect on the RNP was not due to some direct inhibition of NH₃ oxidation metabolism. However, because the eukaryotic growth inhibitors emetine and actinomycin D had no effect on the pasture soil RNP, it remains unclear how cycloheximide is influencing the RNP and needs further investigation.