Research Article

Impaired Reduction of N2O to N2 in Acid Soils Is Due to a Posttranscriptional Interference with the Expression of nosZ

Binbin Liu, Åsa Frostegård, Lars R. Bakken
Mark Bailey, Editor
DOI: 10.1128/mBio.01383-14

ABSTRACT

Accumulating empirical evidence over the last 60 years has shown that the reduction of N2O to N2 is impaired by low soil pH, suggesting that liming of acid soils may reduce N2O emissions. This option has not gained much momentum in global change research, however, possibly due to limited understanding of why low pH interferes with N2O reductase. We hypothesized that the reason is that denitrifying organisms in soils are unable to assemble functional N2O reductase (N2OR) at low pH, as shown to be the case for the model strain Paracoccus denitrificans. We tested this by experiments with bacteria extracted from soils by density gradient centrifugation. The soils were sampled from a long-term liming experiment (soil pH 4.0, 6.1, and 8.0). The cells were incubated (stirred batches, He atmosphere) at pH levels ranging from 5.7 to 7.6, while gas kinetics (NO, N2O, and N2) and abundances of relevant denitrification genes (nirS, nirK, and nosZ) and their transcripts were monitored. Cells from the most acidic soil (pH 4.0) were unable to reduce N2O at any pH. These results warrant a closer inspection of denitrification communities of very acidic soils. Cells from the neutral soils were unable to produce functional N2OR at pH values of ≤6.1, despite significant transcription of the nosZ gene. The N2OR expressed successfully at pH 7.0, however, was functional over the entire pH range tested (5.7 to 7.6). These observations lend strong support to our hypothesis: low soil pH diminishes/prevents reduction of N2O, primarily by precluding a successful assembly of functional N2O reductase.

IMPORTANCE Impaired N2O reduction in acid soils was first observed ~60 years ago, and the phenomenon has been rediscovered several times since then. The practical implication would be that the emissions of N2O from cropped soils could be controlled by soil pH management, but this option has largely been ignored till now. One reason for this could be that the mechanisms involved have remained obscure. Here, we provide compelling evidence that the primary reason is that low pH interferes with the making of the enzyme N2O reductase rather than the function of the enzyme if properly assembled. The implications are important for understanding how pH controls the kinetics of N2O and N2 production by denitrification. The improved understanding provides credibility for soil pH management as a way to mitigate N2O emissions.

INTRODUCTION

Denitrification in soil is the major source of atmospheric N2O (1, 2), which contributes to global warming and destruction of stratospheric ozone. The ultimate driver of the ongoing N2O accumulation in the atmosphere is the input of reactive nitrogen to the biosphere through fertilization and biological nitrogen fixation in agriculture and NOx from combustion (3). This anthropogenic reactive nitrogen will sooner or later return to the atmosphere either as N2O or N2, depending on a plethora of factors which control the N2O/(N2O + N2) product ratio of denitrification within the various parts of the biosphere. This product ratio determines the atmospheric footprint of anthropogenic nitrogen, and the mechanisms controlling this ratio are a key issue where microbial ecology can possibly find solutions to a major environmental problem (4).

Wijler, Delwiche, and Nömmik (5, 6) pioneered investigations of the gaseous products of denitrification in soils and provided the first evidence for a negative effect of acidity on the rate of N2O reduction. Since then, the phenomenon has been rediscovered several times with a variety of soils as summarized by Šimek and Cooper (7), and it now appears an indisputable fact that the N2O/(N2O + N2) product ratio of denitrification in soils tends to increase with increasing acidity of the soils. The implication of this would be that N2O emission from cropped soils could be minimized by management of pH in cropped soils. However, this mitigation option has not gained much momentum, and only a few attempts have been made to rigorously test the effect of pH management on N2O emission in agronomic field experiments as summarized by Qu et al. (8). We believe that there are two main reasons for this lack of interest: the strength of the pH control of the N2O/(N2O + N2) product ratio of denitrification has not been fully appreciated, and the mechanisms involved have not been understood.

Recent experiments with the model organism Paracoccus denitrificans (9) shed some light on the mechanisms. At pH 7, P. denitrificans reduced NO3− to N2 with negligible amounts of N2O emitted. At pH values of <7, transient accumulation of N2O was observed, which increased with decreasing pH, and at pH 6 the cultures produced only N2O. Thus, with decreasing pH, the cells had increasing difficulty in expressing nitrous oxide reductase (N2OR) activity. It was demonstrated that this was not due to a restriction at the transcriptional level. A series of supplementary experiments showed that N2OR, expressed at pH 7, was fully functional at pH 6, although the specific rate was ~50% of that at pH 7. The observed pH response curve of N2O reduction rate was in good agreement with the pH response of N2OR in vitro as determined by Fujita and Dooley (10). The most plausible explanation for these observations is that low pH interferes with the assembly of the enzyme, which takes place in the periplasm (11), where pH is expected to be less controlled than in the cytoplasm (9).

We recently obtained some evidence for a similar posttranscriptional effect of low pH on the expression of N2OR in soil bacterial communities, by measuring gene transcription and rates of N2O reduction in soils from long-term liming experiments (12). In those experiments, where intact soils were investigated, the functional N2OR was produced at substantially lower soil pH than the minimum for this to occur in P. denitrificans. This could suggest that the soils harbor bacteria with a more acid-tolerant expression of N2OR than that of P. denitrificans. Direct evidence for this cannot be provided by such soil incubation experiments, however, due to the potential creation of neutral/alkaline microsites within the soil matrix. This is one reason the pH preference of soil bacteria can only be studied by extracting the cells from the soil matrix (13, 14). Another shortcoming of experiments with intact soil is that it is impossible to test whether functional N2OR, expressed at pH 7, is functional at low pH, as shown for P. denitrificans.

To address these questions, we extracted cells by density gradient centrifugation from soils previously studied (12). The extracted cells were transferred to liquid medium and exposed to a range of pH levels during oxic and subsequently anoxic conditions, while the production/consumption of gases (NO, N2O, and N2) and the transcription of relevant functional genes (nirS, nirK, and nosZ) were monitored. Furthermore, we analyzed the pH robustness of functional N2OR by first allowing the organisms to synthesize the denitrification enzymes at pH 7 and then measuring the potential for N2O reduction at a range of pH levels (5.7 to 7.6).

RESULTS

NO, N2O, and N2 production.To characterize the pH-dependent kinetics of NO, N2O, and N2 production by denitrification, we extracted cells from the three soils (pHs 4.0, 6.1, and 8.0), transferred them to media with 3 pH levels (pHm 5.7, 6.1, and 7.6) in gas-tight serum vials, and replaced the air with helium (He) after a period of 28 h with oxic conditions. The entire procedure from soil to anoxic incubation is illustrated in Fig. 1. At the end of the incubation, measured pH was marginally higher (<0.2 pH units) than the initial pH. The NO, N2O, and N2 production during the anoxic phase is shown in Fig. 2. The pHm had a profound effect on the rate of denitrification and the composition of its gas production. None of the communities were able to produce detectable amounts of N2 at pHm 5.7 and 6.1. At pHm 7.6, the communities from the pHs 6.1 and pHs 8.0 soils produced transient N2O peaks but finally converted all available NO3− to N2 (50 µmol N2 vial−1). The recovery of NO3−–N to gaseous N was close to 100% for treatment, which reached stable plateaus of either N2 or N2O (2 mM NO3− in 50 ml = 100 µmol N).

FIG 1 
FIG 1 

Outline of the oxic/anoxic incubations of cells extracted from soils with different pH (pHs 4.0, 6.1, and 8.0). The cells were incubated in minimal media with different pH (pHm 5.7, 6.1, and 7.6) at 15°C.

FIG 2 
FIG 2 

Kinetics of NO, N2O, and N2 production by bacteria from soils with different pH levels (pHs 4.0, 6.1, and 8.0) during anoxic incubation in medium with three different pH levels (pHm 5.7, 6.1, and 7.6). The figure shows the measured NO (red circle), N2O (green square), and N2 (blue triangle), all as µmol vial−1 (note the different scales). The production of N2 was below the detection limit (~150 nmol vial−1) for all treatments except pHs 6.1 and pHs 8.0 at pHm 7.6. The pH was measured at the end of the incubation and was found to be identical to the initial pH except for a slight increase (<0.2 pH units) in the treatments with high denitrification rates.

In contrast, the community from pHs 4.0 was unable to produce N2 even at pHm 7.6. This lack of N2OR activity was not expected, and we investigated the phenomenon further in a series of experiments at several pHm levels of around 7, using the same procedure as that for the results presented in Fig. 2. This invariably showed that practically all NO3− was converted to N2O. The results of these experiments (including those presented in Fig. 2) are summarized in Fig. 3a, where the N2O indices are plotted against pHm. For the bacteria extracted from pHs 4.0 soil, the indices were around 1.0 at all tested pH levels, i.e., the production of N2 was insignificant.

FIG 3 
FIG 3 

Transient accumulation of N2O and NO by bacteria extracted from soils with different pH (pHs 4.0, 6.1, and 8.0), depending on the pH in the medium (pHm). Panel a shows the N2O index plotted against pHm (each point is the index for a single vial). The index is a measure of the relative amount of N2O, cumulating in equation 1 in the text. Panel b shows the corresponding index for NO (see equation 2 in the text for more details).

Another conspicuous effect of pHm is that the transient NO accumulation declined with increasing pHm. This is clearly seen in Fig. 2 and further elaborated in Fig. 3b, where the NO indices are plotted against pHm (based on the data presented in Fig. 2 and the supplemental experiments with bacteria extracted from the soil with pHs 4.0). In addition to this direct effect of pHm on NO accumulation, the communities were different; NO indices were highest for the community from the most acidic soil.

Since the pHs 6.1 and 8.0 soil communities showed very similar pH-dependent denitrification phenotypes, we decided to concentrate on the soil with intermediate pH (6.1) for testing our core hypothesis regarding the pH dependency of N2O reductase synthesis. This ensured the cells used were extracted from a soil of which the pH was within the experimental pH range (pHm 5.7 to 7.6).

Quantification of functional genes and transcripts.Standard curves for calibration of quantitative PCR were linear for all genes studied (102 to 107 copies; r2 > 0.99, efficiency of ~76 to 87%). Melting curve analysis showed one distinct peak for each expected PCR product and no nonspecific peaks. The primers for nosZ generated a dimer peak in the melting curve analysis, but this did not influence the accuracy of the quantitative PCR since the data were collected at 82°C, where the primer dimer had been denatured and no fluorescent signals were emitted.

Figure 4 shows the copy numbers of nirK, nirS, and nosZ genes (a to c) and transcripts (d to f) throughout the anoxic incubation of cells from pHs 6.1 at pHm 5.7, 6.1, and 7.6. The copy number of all genes increased sharply during the first 20 h of anoxic incubation at pHm 7.6. At pHm 6.1, a similar increase was observed for the nosZ and nirS genes (although they reached lower levels than at pHm 7.6), whereas the copy numbers of nirK remained practically constant. At pHm 5.7, the copy numbers of nosZ and nirS showed a slow increase throughout the incubation. The initial copy numbers of nirK genes declined with pH, reflecting changes during the 28 h of oxic preincubation at the respective pHm (see Fig. 1). Although pHm clearly affected the growth of denitrifying bacteria, both during the 28-h oxic preincubation (resulting in different initial number of gene copies during the anoxic incubation) and during the anoxic incubation, the pHm had no consistent effect on the ratio between the copy numbers of nosZ and nir (nirK + nirS), as shown in Fig. S1 in the supplemental material.

Figure S1

Ratios of the functional genes nos vs. nir and ratios of the transcripts of the same genes, measured in cells extracted from soil samples with pH 6.1 (pHs 6.1) and incubated for 100 h with different pH levels in the medium (pHm 7.6, 6.1, and 5.7). Download Figure S1, DOCX file, 0.1 MB.
Copyright © 2014 Liu et al.

This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported license, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

FIG 4 
FIG 4 

Copy numbers of the nirS, nirK, and nosZ genes (a to c) and the corresponding transcripts (d to f) throughout the 100 h of anoxic incubation of cells extracted from soil in which the pH (pHs) was 6.1 and incubated in media of different pH (pHm 5.7, 6.1, and 7.6) (see Fig. 1). Results are shown in a separate plot for each pH level of the medium. Quantitative PCR was performed in triplicates for each sample, and average values are shown (average coefficients of variation were 32% for DNA and 40% for cDNA). Time = 0 is the time of oxygen removal.

Transcription of the nirS, nirK, and nosZ genes was quantified for the pHs 6.1 samples when incubated in media with pHm 5.7, 6.1, and 7.6. For nirS and nosZ genes, the transcription, presented as mRNA copy numbers, increased sharply during the first 10 to 15 h and reached somewhat higher levels at pHm 7.6 than at pHm 6.1 (Fig. 4). The nirK mRNA copy numbers were 1 to 2 orders of magnitude lower than those of nirS.

To evaluate whether pHm had any effect on the transcriptional regulation of the three genes, we calculated the number of transcripts per gene (i.e., cDNA/DNA). Figure 5 shows the average values (cDNA/DNA) for the first 40 h of incubation (the ratios throughout the anoxic incubation are shown in Fig. S2 in the supplemental material). For nosZ, these average transcription ratios were low (0.01 to 0.02) and essentially independent of pHm. Higher values were recorded for nirS, with a trend toward lower values at the highest pH, although this was not statistically significant (P = 0.33, one-way analysis of variance [ANOVA]). This trend was even more pronounced for nirK (and statistically significant, P = 0.007).

Figure S2

Ratio between copy number of transcripts and copy number of genes, plotted against time. Results are shown as separate plots for each gene and separate graphs within plots for each pH level in the medium (pHm 5.6, 6.1, and 7.6). Download Figure S2, DOCX file, 0.2 MB.
Copyright © 2014 Liu et al.

This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported license, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

FIG 5 
FIG 5 

Average transcription ratios for nirS and nosZ genes (mRNA copies/DNA copies) during the first 40 h of anoxic incubation. The values are based on data presented in Fig. 4. Standard deviations are shown as vertical lines.

Since the gene copy numbers increased substantially during the first 20 h of anoxic incubation at pHm 6.1 and 7.6 (Fig. 4), one might suspect a major shift in the bacterial community composition due to growth of one or a few populations, representing only a small fraction of the originally extracted bacteria. We evaluated this qualitatively by running denaturing gradient gel electrophoresis (DGGE) analyses of the nosZ genes during the first 80 h of the pHm 7.6 incubation of cells extracted from soil with pHs 6.1. The results show a large number of bands which were essentially unaltered during the whole incubation period (see Fig. S3 in the supplemental material).

Figure S3

DNA DGGE fingerprinting of the nosZ gene from the extracted cells (from soil pHs 6.1) during anoxic incubation at pHm 7.6 (gas kinetics shown in Fig. 2). Lanes 1 to 5 are samples taken at time 0 h, 20 h, 40 h, 60 h, and 80 h, respectively. Download Figure S3, DOCX file, 0.1 MB.
Copyright © 2014 Liu et al.

This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported license, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

The cells extracted from the pHs 4.0 soil were unable to make functional N2OR even at pH 7.0 to 7.6 (Fig. 2 and 3). To investigate if this could be due to lack of nosZ transcription, gene transcripts for this community were quantified during incubation in medium with pHm 7.6. The results showed that the copy numbers of nirS and nosZ transcripts were in the same range as for cells extracted from pHs 6.1 soil, but the cells were unable to reduce N2O to N2 (Fig. 6).

FIG 6 
FIG 6 

Transcription of nirS and nosZ during denitrification at pHm 7.6, by bacteria extracted from soil with pHs 4.0. The number of transcripts per vial is shown on the left y axis, and the accumulated N2O production is shown on the right y axis. N2 production was below the detection limit during the entire incubation.

pH sensitivity of N2OR.The direct effect of pH on N2OR activity and oxic respiration was tested by first producing cells with an intact denitrification proteome developed at pH 7.0 (weakly buffered medium), which were then used to inoculate vials with strongly buffered medium covering a pH range from 5.7 to 7.6 (lower pH levels could be desirable but would require another buffer system). Two sets of vials were incubated, both with a medium without NO3− and NO2− but with different treatments regarding headspace gas composition. Set 1 had 10 ml O2 liter−1 in the headspace, in order to measure oxic respiration. Set 2 had anoxic (He) headspace with N2O (~5.4 ml liter−1) and nitrate-free medium, in order to measure the potential rate of N2O reduction. The outline of the entire experiment is illustrated in Fig. S4 in the supplemental material. The results are summarized in Fig. 7, where the average rates during the first 5 h of incubation are plotted against pHm. The oxic respiration rate (Fig. 7A) was ~1 µmol O2 vial−1 h−1 and not much affected by pHm. The rates of N2O reduction (Fig. 7B) were ~1.15 µmol N2O vial−1 h−1 at pH 7.2 and 7.6 and fell gradually with pH to reach ~0.6 µmol N2O vial−1 h−1 at pH 5.7.

Figure S4

Outline of the experimental procedures for testing the direct effect of pH on N2OR expressed at pHm 7.0, where pHm indicates the pH of the medium. Download Figure S4, DOCX file, 0.1 MB.
Copyright © 2014 Liu et al.

This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported license, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

FIG 7 
FIG 7 

Effect of pHm (pH level of the medium) on the metabolism of a denitrifying community with functional N2O reductase was expressed at pH 7.0. The cells (extracted from soil with pH 6.1) were allowed to express denitrification at pH 7.0 and then transferred to vials with pHm 5.7 to 7.6. (A) Rate of oxic respiration measured in vials with oxic headspace; (B) rate of N2O reduction in vials with anoxic headspace containing N2O (and NO3−-free medium). All rates are averages from the first 5 h of incubation (i.e., after transfer to the different pHm levels), and standard deviations are shown as vertical bars (3 replicate vials for panel A, n = 6 for panel B).

DISCUSSION

The primary denitrification experiments (Fig. 1 and 2) were designed to investigate whether any of the soils along the pH gradient from 4.0 to 8.0 harbored denitrifying bacteria that could produce functional N2OR at lower pH than the limit observed for P. denitrificans (9). The results indicate that this is not the case; all three soil communities were unable to produce significant amounts of functional N2OR at pHm values of ≤6.1 within the time span of the incubation experiments (~100 h), suggesting that the published results from the denitrification model organism demonstrate a more general phenomenon applicable to a wide range of different bacteria. This leads to the obvious question of how organisms in intact soils with pH values of ≤6.1 were indeed able to reduce some N2O to N2 (albeit later/slower than in the more alkaline soils) under anoxic conditions, as demonstrated in our previous study (12). Our tentative explanation is that the successful making of N2OR in moderately acid soils (i.e., pH of ~6.0) takes place in neutral/alkaline microsites with higher pH than the average (bulk) pH of the soil. Such microsites may be created within and around clusters of actively denitrifying cells. This has been demonstrated in biofilms (15) and utilized to differentiate between denitrifying and nondenitrifying colonies in agar (16). We are not aware of any studies of this phenomenon in soils, but alkalinization (1 to 2 pH units) of the soil close to roots during active uptake of NO3− has been demonstrated (17). If the finding for the model organism P. denitrificans (9) also applies to other bacteria, a local pH value of >6.1 is needed to make the N2OR. This would be increasingly difficult with increasing acidity of the soil, thus explaining the observation that the transient peak of N2O increased gradually with increasing soil acidity (12). An interesting extrapolation of this is that the distribution of denitrification activity within the soil matrix (clumped in large colonies versus dispersed throughout the soil) could have an effect on the N2O emission: locally neutral/high pH may be reached in microsites with a high density of actively denitrifying bacteria (hence production of functional N2OR), whereas single cells within the soil matrix are unlikely to achieve this.

We were also interested in the kinetics of NO accumulation during anoxic respiration since NO is considered a strong inducer of transcription of nir and nor, coding for nitrite and nitric oxide reductase, respectively (18). NO was also recently found to induce nosZ transcription in P. denitrificans (19). The results (Fig. 2) demonstrate that the transient accumulation of NO decreased with increasing pHm and that the community from the most acidic soil produced more NO than those from the more alkaline soils compared at the same pHm (Fig. 3). Thus, low pH soils appear to select for organisms that produce large amounts of NO due to imbalanced denitrification. Imbalanced denitrification leading to a high net production of NO was recently demonstrated for Agrobacterium tumefaciens in response to changing oxygen levels (20, 21). An alternative explanation could be that low-pH soil selects for organisms that reduce all nitrate to nitrite prior to the reduction of nitrite, which has been observed for certain strains within the genus Thauera (22). Nitrite accumulation would enhance NO emission at moderately low pH due to chemical decomposition (23).

By quantification of gene transcription as a function of pHm, we wanted to clarify if pH affected the transcriptional regulation of nosZ. The data as presented in Fig. 4 (copies vial−1) could suggest that the transcription of nosZ is strongly repressed by low pH. However, transcription must be evaluated against the copy number of the same genes. The copy number of nosZ genes was clearly higher in the vials with higher pHm, even at the onset of the anoxic incubation. This probably reflects that the organisms harboring the nosZ genes grow faster at high than at low pH, both during the oxic incubation and during the subsequent anoxic incubation. A stringent test is thus to calculate the ratio of mRNA copies/DNA copies of nosZ as shown in Fig. 5. This suggests that pH had no effect on the transcriptional regulation of the nosZ genes (the mRNA/DNA ratio for nosZ was not much affected by pH). It should be kept in mind, though, that PCR-based quantifications of genes and transcripts are biased. The PCR primers used in this study will not capture all the genes coding for N2OR (24). However, the mRNA/DNA ratios should be valid for the genes amplified with the primers used, since cDNA and DNA were amplified by the same set of primers. Thus, for the organisms harboring these genes, there is no evidence for a pH-dependent transcription of the nosZ gene. The result corroborates earlier findings (9, 12) that transcriptional regulation provides no explanation for the low N2OR activity in acid soils.

The results for the cells extracted from soil with pH 4.0 are difficult to understand for several reasons. One would expect that if denitrifying bacteria with an acid-tolerant N2OR expression exist, they should be selected in the most acidic soil, but the results lend no support to this notion; the bacteria extracted from the most acidic soil were unable to produce functional N2OR even at pH values around 7.0 (Fig. 2 and 3). We hypothesized that the complete lack of N2OR activity in the pHs 4.0 cells could be ascribed to a “silencing” of the nosZ genes in this soil through mutation of the nosZ gene itself or another gene necessary for its expression, such as nosR (25). The latter was reported in a recent, comparative study of denitrification in strains within the genus Thauera, of which one strain was unable to reduce N2O to N2, apparently due to the absence of an intact nosR gene (22). However, the quantification of transcripts (Fig. 6) in cell extracts from the pHs 4.0 soil in the present study lends no support to such lack of transcription, at least for the organisms whose nosZ genes were captured by our primers. An alternative explanation is that mutations may have occurred in other genes coding for factors involved in nosZ maturation or, as discussed above, that those factors do not function at low pH.

The complete lack of N2 production by the cells from the soil with pHs 4.0, even at pHm values of ≥7, is in stark contrast to recent findings in our laboratory that this soil is able to express N2OR quite effectively in response to liming (unpublished data). This may suggest a serious bias in the cell extraction by density gradient centrifugation. Nadeem et al. (26) recently investigated this by comparing the kinetics of N2O and N2 production by loosely attached and strongly attached cells (the latter extracted by vigorous dispersion of the pellet beneath the density gradient of the first extraction). The results demonstrated a moderate bias in the sense that the loosely attached cells produced less N2 and more N2O than the strongly attached cells. This bias might be more severe for the soil with pH 4.0 than for the other soils, thus reconciling the apparently conflicting observations of the response to increased pH (intact soil responding by N2O reduction, extracted cells not). This does not resolve the puzzle, however, that the cells extracted from this soil were notoriously unable to reduce N2O to N2 despite significant transcription of nosZ (Fig. 6).

In the last set of experiments, cells with an intact denitrification proteome, expressed at pH 7.0, were challenged to reduce N2O at a range of pH values (Fig. 7). The results strongly support our hypothesis that the successfully assembled N2OR is functional over a wide pH range and provide indirect evidence that the complete inability of the indigenous soil bacteria to produce active N2OR at pH values of ≤6.1 is a posttranscriptional problem which is due to interference with the making of the enzyme. At present, we do not know why organisms are unable to make functional N2OR at pH values of ≤6.1. Based on the results with P. denitrificans, we hypothesized that it may be due to interference with the assembly of the protein in the periplasm where the insertion of Cu2+ is possibly hindered by low pH (9). If so, the enzyme could be “repaired” by increasing the pH, as demonstrated for copper-deficient N2OR in Pseudomonas stutzeri (27). In theory, this may even happen at low pH, albeit so slowly that it will not be observed within the time frame of our incubation experiments. Further investigations are needed to clarify these questions.

One would expect that acid environments select for organisms which are able to produce active N2OR successfully at low pH. Our data provide compelling evidence that such organisms are not present in the soils studied. Is there any evidence for it elsewhere? Palmer and Horn (28) found high rates of N2O reduction in acid (pH 4.4) peat soils, determined by anoxic incubation of soil slurries. The N2O reduction was ascribed to denitrification driven by organisms with nirS rather than nirK. This is hardly evidence for acid-tolerant production of N2OR, since there may exist neutral/alkaline hot spots within the soil matrix, as discussed above. Green et al. (29, 30) studied denitrifying communities in contaminated subsurface soils and found that environments with low pH (3, 4) selected for denitrifying bacteria within the genus Rhodanobacter. They concluded that these organisms represent an acid-tolerant group of denitrifiers, able to reduce NO3− all the way to N2. Six isolates were recently genome sequenced, confirming the presence of nirK and nosZ genes, although three of the isolates lacked the gene for nitrate reductase (31). To our knowledge, no stringent test has been conducted to evaluate the ability of Rhodanobacter strains to express N2OR at low pH. However, the investigation by van den Heuvel et al. (32) sheds some light on this. They grew denitrifying communities from acid soils in continuous reactors at variable pH and found that low pH selected for Rhodanobacter strains which were evidently able to denitrify at pH as low as 4. However, although the communities were able to reduce NO3− to N2 when grown at neutral pH, they produced practically pure N2O at pH values of <6.2. Thus, it may well be that Rhodanobacter strains are adapted to denitrify at low pH, but they appear to have the same difficulties with producing active N2OR at such low pH as other bacteria.

We are aware that the primers used for nosZ in this study fail to amplify atypical nosZ genes claimed to be important in soils (33). However, this would hardly affect the validity of our conclusions regarding the lack of functional N2O reductase at low pH and the robust functioning of N2O reductase (expressed at pH 7.0) at pH below 6. We are also aware that the composition of the microbial communities may have an impact on denitrification rates, the gaseous product composition, and the emissions from soils as explored in several studies, which are reviewed recently by Braker and Conrad (34). One common notion in these studies has been that high N2O emission could be ascribed to the absence of organisms carrying nosZ (35). This appears irrelevant in our case, however, since the nosZ gene pool was abundant at all 3 pHm levels (12).

In conclusion, we are beginning to understand why soil acidity exerts a pervasive control of the N2O/(N2 + N2O) product ratio of denitrification and hence on the propensity of soils to emit N2O. Some empirical evidence for a direct effect of pH on N2O emissions has been provided by studying the spatial distribution of N2O emission and pH in forested riparian soils (36) and grassland soils (37). More rigorous testing is clearly needed to evaluate this. The immediate effect of liming acid soils may be to enhance N2O emissions; however, as demonstrated by Clough et al. (38) and Baggs et al. (39), the reason for this is that liming acid soils will induce a transient enhancement of C and N mineralization and nitrification and hence also the rate of denitrification. The net outcome may thus be that liming induces a transient enhancement of N2O emission despite a lowered N2O/(N2 + N2O) product ratio of denitrification. However, the long-term effect of sustaining a high pH in agricultural soils is most likely to secure low N2O emissions.

MATERIALS AND METHODS

Soil and extraction of bacteria.Peat soils (Sapric Histosol, FAO/ISRIC/ISS, containing 45% organic C, 2% organic N) were sampled from a soil liming experiment (pHs 4.0 to 8.0) established in 1978 (40), which was used previously for investigating N2O production by nitrification (41) and denitrification (12). The soils were sieved (6 mm) while moist and stored moist (1.5 ml water g−1 soil [dry weight], i.e., ~50% air-filled porosity) at 4°C until used. The outline of the preincubation, cell extraction, and subsequent incubations is shown in Fig. 1. To raise the microbial biomass/activity prior to cell extraction, moist soil samples (20 g fresh weight = 8 g dry weight) were mixed with finely ground clover leaves (5 mg/g soil [dry weight]) and incubated for 85 h at 15°C (monitored for respiration rate and NO and N2O production [see Fig. S5 in the supplemental material]). Cells were then extracted by dispersion/density gradient centrifugation as described by Bakken and Lindahl (42). The pH of soil slurries of pH 6.1 and 4.0 soils was adjusted to 7.0 prior to dispersion. The total number of cells extracted was determined by epifluorescence microscopic counts (43) in preliminary experiments (7 × 109 to 9 × 109 cells g−1 soil [dry weight], which is 10 to 12% of the total numbers of cells in the soils; no statistical difference between soils as tested by one-way analysis of variance [ANOVA]).

Figure S5

CO2 production and N2O and NO kinetics of intact soils with pHs 4.0, 6.1, and 8.0 in response to clover amendments. Soil samples (20 g) were incubated in gas-tight 120-ml medical vials, and gases were monitored during 85 h. (A) Accumulated CO2 production (μmol CO2 g−1 soil); (B) measured amount of N2O (nmol vial−1) and pH 4.0 soil amended with clover are shown on the right y axis; (C) measured amount of NO (nmol vial−1) and pH 4.0 soil amended with clover are shown on the right y axis. Download Figure S5, DOCX file, 0.5 MB.
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Culture conditions and general incubation procedure.The medium was the same as that used by Morley et al. (44): 3 mM (NH4)2SO4, 5 mM glutamic acid, 1.76 mg liter−1 EDTA, 10 mg liter−1 ZnSO4, 5 mg liter−1 FeSO4, 1.5 mg liter−1 MnSO4, 0.4 mg liter−1 CuSO4, 0.25 mg liter−1 Co(NO3)2, 0.15 mg liter−1 H3BO3, 1 mg liter−1 nicotinic acid, 0.5 mg liter−1 thiamine, and 1 mg liter−1 biotin, buffered with sodium phosphate (10 mM for the oxic preculturing and 100 mM for the denitrification experiments). The concentration of NO3− was either 1 or 2 mM (KNO3), except when determining potential N2O reduction, for which NO3−-free medium was used, prepared as described by Bergaust et al. (19).

All culturing was done in serum vials (120 ml) with butyl rubber septa, containing a 50-ml culture volume, vigorously stirred by Teflon-coated magnetic bars and placed in a robotized incubation system (at 15°C) which monitors the headspace concentrations of O2, CO2, NO, N2O, and N2. The system is described in detail by Molstad et al. (45), who also describe the techniques to replace headspace atmosphere with He prior to inoculation.

The extracted cells (~10 ml) were transferred to vials containing 40 ml weakly buffered medium (10 mM phosphate, pH 7.0) and incubated (oxic) for 8 h. The precultured suspensions were then used to inoculate vials (5 ml vial−1) containing 45 ml strongly buffered medium (100 mM phosphate, 2 mM KNO3) at three different pH levels (pHm 5.7, 6.1, and 7.6), which were first incubated for 28 h under oxic conditions (oxic incubation [28 h], Fig. 1) while respiration rates were monitored. Measured respiration and apparent oxic growth rates (0.14 to 0.25 h−1) are shown in Fig. S6 and S7 in the supplemental material. The motivation for these oxic preincubations was to secure a low level of intact denitrification enzymes in the cells prior to the anoxic incubation (preexisting denitrification enzymes would be diluted by oxic growth).

Figure S6

Oxygen consumption during the oxic phase of extracted cell incubations (prior to anaerobization; see Fig. 1). The graphs show the accumulated amounts of oxygen consumed (i.e., integrals of measured rates), plotted against time. The results for the three soils are shown in separate panels (pHs 8.0, 6.1, and 4.0). Within each panel, the results for the different media (pHm 5.7, 6.1, and 7.6) are shown as separate curves (single vial results are shown; two replicate vials were monitored for the pHm 6.1 and pHm 7.6 treatments). Note that the scale is logarithmic. Download Figure S6, DOCX file, 0.1 MB.
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The vials were then made anoxic by evacuating and filling with pure helium (He) twice (reducing oxygen concentration to ≤250 µl liter−1 = 0.4 µM in the liquid), and the incubation was continued for 100 h (anoxic incubation, Fig. 1). The incubation robot only takes 15 vials, thus it was not possible to include replicates for all 9 treatments. However, the experiment was repeated several times (n ≥ 3), both for the community extracted from the soil with pHs 6.1 (for quantification of genes and transcripts, see below) and for the community extracted from the most acidic soil (pHs 4.0). There was generally little variation in gas kinetics results between replicates within and between experiments, as seen in Fig. 3, which condenses the results in one graph.

The NO and N2O indices (12) were calculated in order to summarize the effect of pHm on the transient accumulation of NO and N2O as a fraction of the total N gas produced: IΝ2Ο=∫0TΝ2Ο(t)/[∫0TΝ2Ο(t)+∫0TΝΟ(t)+∫0TΝ2(t)](1) IΝΟ=∫0TΝΟ(t)/[∫0TΝ2Ο(t)+∫0TΝΟ(t)+∫0TΝ2(t)](2) where NO(t), N2O(t), and N2(t) are the amounts of the three gases at any time t and T is the time when all NO3− is recovered as gas (NO + N2O + N2).

Determination of the effect of low pH on denitrification enzymes expressed at high pH.To determine the direct effect of pH on the activity of the denitrification enzymes, cells extracted from soil with pHs 6.1 were incubated at pHm 7.0 (10 mM phosphate buffer) under anoxic conditions and allowed to deplete the 2 mM NO3− in the medium (recovered as N2 in the headspace). This culture was then used to inoculate a series of vials containing 45 ml strongly buffered medium (100 mM phosphate buffer) with five different pHm levels ranging from 5.7 to 7.6. We prepared two sets of vials for these experiments: (i) the first set had oxic headspace (10 ml O2 liter−1) and was used to determine the oxic respiration; (ii) the second set had NO3−-free medium and He atmosphere to which we added ~0.6 ml pure N2O, resulting in ~5.4 ml N2O liter−1 in the headspace (~170 µM N2O in the liquid). These vials were used to obtain a direct measurement of the rate of N2O reduction.

For both sets, O2, N2O, and N2 concentrations were monitored by sampling every 40 to 60 min for a period of 20 h. The measured rates (i.e., O2 consumption in set 1; N2O reduction in set 2) were fairly constant throughout the first 5 to 7 h of incubation, as illustrated for set 2 (see Fig. S8 in the supplemental material). Since the purpose of these experiments was to determine the direct effect of the change in pH, we report only the average rates during the first 5 h of incubation.

Figure S8

Measurement of N2O reduction rates and N2 production rates. Actively denitrifying cultures (extracted cells from pHs 6.1 soil, grown at low buffer concentrations, pHm 7) which had depleted the NO3− were injected into vials with NO3−-free medium (with high buffer concentrations) which contained ~5 ml N2O liter−1 in the headspace. The example shown is a single-vial result with pHm 7.6. The top panel shows the measured amounts of N2O and N2 (headspace + liquid). The second panel shows the calculated rates of N2O reduction and N2 production for each time increment. The last panel shows the cumulative reduction of N2O and production of N2. Regression functions for accumulated N2O and N2 based on data for the first 7 h are shown. The reason for the decline in N2 after 14 h is sampling loss (3.4% of headspace is removed and replaced by He for each sampling). These sampling losses are taken into account when calculating the rates (both for N2 and N2O). Download Figure S8, DOCX file, 0.1 MB.
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Extraction of nucleic acids.Gene expression was measured on cells from the pHs 6.1 soil incubated at pHm 5.6, 6.1, and 7.6 in a separate water bath, in parallel with the experiment in the incubation robot (stirring and temperature conditions were identical). Three replicate vials were sampled (entire vials) at each time point shown in Fig. 4. The cells were collected by centrifugation (10,000 × g, 10 min at 4°C) and resuspended in 1 ml of RNAprotect (Qiagen, Nordic-Norway) and quickly frozen at −20°C. The same treatment was also performed for cells form pHs 4.0 soil incubated at pHm 7.6 medium.

Total nucleic acids were isolated using a bead-beating method. To avoid RNase contamination, only certified nuclease-free tubes and diethyl pyrocarbonate (DEPC)-treated solutions (46) were used. The cell lysis, protein denaturation, and nucleic acid purification were performed by a method described in reference 47 with modifications (12, 48). The nucleic acid extracts were split into two portions, and these were treated with RNase and DNase, respectively. The extracts intended to contain only RNA were treated with an RNase-free DNase set (Qiagen, Nordic-Norway) for 30 min, but this did not remove all genomic DNA. An additional 2.5-h treatment was conducted with the DNase I kit (Sigma-Aldrich). This resulted in successful removal of all genomic DNA, as seen from the absence of PCR products from control samples that had been treated with DNase but not with reverse transcriptase. The A260/A280 ratios of both DNA and RNA were higher than 1.7 for all samples.

Reverse transcription.Reverse transcription (RT) was performed using the Superscript RT-PCR system (Invitrogen) with the method described in the manual. Two master mix solutions were prepared in separate 0.2-ml tubes. Master mix 1 contained 1 µl deoxynucleoside triphosphate (dNTP) mix (10 mM each; TaKaRa Bio Co., Japan), 2 µl random hexamer primers (50 ng/µl; Promega), total RNA, and RNase-free water up to 13 µl. Master mix 2 contained 4 µl 5× first-strand buffer, 1 µl 0.1 M dithiothreitol (DTT), 1 µl Superscript III RT enzyme (200 units/µl), 1 µl RNaseOUT recombinant RNase inhibitor (Invitrogen; 40 units/µl). Master mix 1 solution was incubated at 65°C for 5 min and put on ice for 2 min. The contents of the tube were collected by brief centrifugation before master mix 2 was added. The tube was transferred to a thermocycler (Applied Biosystems) and incubated at 25°C for 5 min, 50°C for 60 min, and 70°C for 15 min. The cDNA was stored at −20°C until quantitative PCR analysis. Control samples containing the same concentration of RNA, but on which reverse transcription was not performed, were prepared for each gene (nirS, nirK, and nosZ) and subjected to quantitative PCR to ensure that all genomic DNA was removed by DNase.

Quantitative PCR.The genes and their transcripts were quantified with a StepOne Plus quantitative PCR system (Applied Biosystems). Gene fragments of nirS, nirK, and nosZ were amplified with the primer pairs cd3af, r3cd (49, 50), 1F, 5R (51), Z-F, 1622R (50, 52), respectively. A standard curve method was employed, since previous studies using the same experimental setup for batch culture incubation showed that the use of a housekeeping gene as the endogenous standard generated unstable expression levels due to the rapid changes in bacterial activity and growth (20). Construction of the standard curves, as well as the quantitative PCR program, was as described previously (12).

DGGE.DGGE was performed on extracts from pHs 6.1 soil, sampled during the incubation shown in Fig. 2. The samples from pHm 7.6 medium were taken during the anoxic incubation period at 0 h, 20 h, 40 h, 60 h, and 80 h. The nosZ gene was amplified with primer pair ZF and 1622R-GC (50, 52). DGGE of PCR products was performed using the method described by Muyzer et al. (53) with the use of a Dcode system (Bio-Rad Laboratories). Polyacrylamide gradient gel was made by 8% polyacrylamide with 35 to 60% denaturant (100% defined as 7 M urea and 40% formamide). The gel was run for 200 min at 200 V in 1× Tris-acetate-EDTA (TAE) buffer at a constant temperature of 60°C. The gels were stained with ethidium bromide and photographed in a GelDoc XR UV light imager (Bio-Rad).

Statistical analyses.Statistical analyses were performed using Minitab 16 (Minitab Inc., State College, PA, USA). One-way ANOVA was used to compare the cell extraction efficiencies for the different soils and for testing if the pH in the media affected the mRNA/DNA ratio for the three genes in question. Preliminary analysis of the mRNA/DNA ratio showed that the distribution of the residuals deviated significantly from normal distribution (P < 0.01; tested by Kolmogorov-Smirnov’s test for normality). Hence, we analyzed the log-transformed data, for which the distribution of residuals did not significantly deviate from normal distribution (P > 0.15). For cell extractions, log transformations prior to ANOVA were not necessary (the distribution of residuals did not deviate significantly from the normal distribution).

Figure S7

Apparent growth rates for cells extracted from soils with different pH (pHs 4.0, 6.1, and 8.0) during the oxic phase prior to anaerobization. The apparent growth rates (h−1) were estimated by regression [linear regression of ln(Vresp) against time, where Vresp is the measured respiration rate during the 28-h oxic preincubation]. Results are shown for each community (i.e., cells extracted from soils with three different pH levels and incubated in medium with pHm 5.7, 6.1, and 7.6), plotted against the pHm. Download Figure S7, DOCX file, 0.1 MB.
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FOOTNOTES

    • Received 29 May 2014
    • Accepted 2 June 2014
    • Published 24 June 2014

This is an open-access article distributed under the terms of the Creative Commons Attribution-Noncommercial-ShareAlike 3.0 Unported license, which permits unrestricted noncommercial use, distribution, and reproduction in any medium, provided the original author and source are credited.

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