Editor: Christoph Tebbe
Abundance, diversity and functional gene expression of denitrifier communities in adjacent riparian and agricultural zones
Article first published online: 25 MAR 2011
© 2011 Federation of European Microbiological Societies. Published by Blackwell Publishing Ltd. All rights reserved
FEMS Microbiology Ecology
Volume 77, Issue 1, pages 69–82, July 2011
How to Cite
Dandie, C. E., Wertz, S., Leclair, C. L., Goyer, C., Burton, D. L., Patten, C. L., Zebarth, B. J. and Trevors, J. T. (2011), Abundance, diversity and functional gene expression of denitrifier communities in adjacent riparian and agricultural zones. FEMS Microbiology Ecology, 77: 69–82. doi: 10.1111/j.1574-6941.2011.01084.x
Present address: Catherine E. Dandie, Adelaide Laboratory, CSIRO Land and Water, PMB 2, Glen Osmond, SA 5064, Australia.
- Issue published online: 6 JUN 2011
- Article first published online: 25 MAR 2011
- Accepted manuscript online: 8 MAR 2011 10:01AM EST
- Received 22 September 2010; revised 18 February 2011; accepted 3 March 2011., Final version published online 25 March 2011.
- nitrous oxide;
- slope position;
- quantitative PCR
- Top of page
- Materials and methods
- Supporting Information
Lands under riparian and agricultural management differ in soil properties, water content, plant species and nutrient content and are therefore expected to influence denitrifier communities, denitrification and nitrous oxide (N2O) emissions. Denitrifier community abundance, denitrifier community structure, denitrification gene expression and activity were quantified on three dates in a maize field and adjacent riparian zone. N2O emissions were greater in the agricultural zone, whereas complete denitrification to N2 was greater in the riparian zone. In general, the targeted denitrifier community abundance did not change between agricultural and riparian zones. However, nosZ gene expression was greater in the riparian zone than the agricultural zone. The community structure of nirS-gene-bearing denitrifiers differed in June only, whereas the nirK-gene-bearing community structure differed significantly between the riparian and the agricultural zones at all dates. The nirK-gene-bearing community structure was correlated with soil pH, while no significant correlations were found between nirS-gene-bearing community structure and soil environmental variables or N2O emissions, denitrification or denitrifier enzyme activity. The results suggested for the nirK and nirS-gene-bearing communities different factors control abundance vs. community structure. The nirK-gene-bearing community structure was also more responsive than the nirS-gene-bearing community structure to change between the two ecosystems.
- Top of page
- Materials and methods
- Supporting Information
Riparian buffer zones are an excellent management practice for the protection of aquatic ecosystems from nutrient runoff from agricultural fields (Mayer et al., 2007). Denitrification is one metabolic pathway that removes nitrogen (N) in riparian zones (Verhoeven et al., 2006; Woodward et al., 2009). Denitrification, a microbial process, reduces nitrate (NO3−) and nitrite (NO2−) to the gases nitric oxide (NO), nitrous oxide (N2O) and dinitrogen (N2). N2O can be emitted as a consequence of incomplete denitrification (Knowles, 1982). N2O is a greenhouse gas, with a global warming potential ∼298 times that of carbon dioxide (CO2) (IPCC, 2007). N2O accounts for 7.9% (CO2 equivalent emission) of anthropogenic greenhouse gas emissions and atmospheric concentrations of N2O have been increasing since preindustrial times at a rate of about 0.26% per year (Forster et al., 2007).
Riparian and agricultural zones differ in soil properties, soil water content, plant species and nutrient concentration; hence, denitrification rates and N2O emissions may be regulated by different factors in each environment. Riparian zones generally have high concentrations of organic carbon (C) and high soil water content, resulting in the formation of anoxic conditions that are conducive to complete denitrification (Davis et al., 2008). Through the application of N fertilizer, agricultural soils often contain elevated NO3− concentrations and previous studies have reported high rates of denitrification and N2O emissions in agricultural soils, especially in humid environments (Bateman & Baggs, 2005; Hofstra & Bouwman, 2005). The distinct differences in the environmental conditions between agricultural and riparian zones may influence denitrifier community dynamics and may also directly influence denitrifier activity, and therefore, denitrification and N2O emissions. Only one study to date, to the best of our knowledge, has investigated denitrifier community structure in cropped fields and adjoining riparian areas (Rich & Myrold, 2004). In that study, the community structure of nosZ-gene-bearing denitrifiers was studied using terminal-restriction fragment length polymorphisms and distinct nosZ gene community structures were observed between riparian and agricultural zones.
Denitrifiers are a critical functional guild in the N cycle. Functional genes (nirK, nirS, cnorB, qnorB, nosZ) involved in the denitrification pathway have been used as genetic markers for denitrifier abundance, diversity and more recently for functional gene expression (as reviewed in Philippot et al., 2007). There is still a limited understanding of the controls on denitrifier abundance and diversity in the environment and of the relationships between denitrification activity and denitrifier community structure and/or abundance (Enwall & Hallin, 2009; Philippot et al., 2009). Some studies have shown relationships between community structure and denitrification activity (Wertz et al., 2009; Enwall et al., 2010) and denitrifier abundance and potential denitrification activity (Hallin et al., 2009; Philippot et al., 2009; Enwall et al., 2010). However, other studies reported that denitrifier community structure (Rich & Myrold, 2004; Boyle et al., 2006) or denitrifier abundance and denitrification gene expression (Dandie et al., 2008; Henderson et al., 2010) were uncoupled from variations in the denitrification rate and N2O emissions in soil.
The aim of this study was to characterize the differences in the abundance, diversity and denitrification gene expression of denitrifying communities between an agricultural zone cropped to maize (Zea mays L.) and a riparian zone (mixed wood with a herbaceous understorey; Leclair, 2010) adjacent to Thomas Brook, NS, Canada (N4504.886 W6445.318). The agricultural–riparian zone delineation has been present at this site for over 100 years. This study is part of a larger 2-year study looking at the effect of landscape position and land-use zone on greenhouse gas fluxes (i.e. N2O, CO2, CH4; Leclair, 2010), while the current study focused on land-use effect on denitrifier communities on three dates in 2007. Sampling at the site was designed to study the effect of slope position and land use on denitrifying communities. Denitrifying communities were monitored based on functional genes for enzymes in the denitrification pathway (nirK, nirS, nosZ). Secondary to this aim, the differences in denitrifying communities were compared with parameters related to denitrification (potential denitrification enzyme activity (DEA) assay, denitrification rate and N2O flux) and other soil physicochemical properties (i.e. pH, NO3− concentration and others) to determine the linkages between functional gene-bearing communities and function or environmental parameters in these adjacent ecosystems. We hypothesized that (1) soil and environmental properties vary among slope positions and between land-use zones; (2) these differences in soil and environmental factors influence denitrifier community abundance and structure; and (3) these changes in denitrifier community abundance and structure result in differences in denitrification activity.
Materials and methods
- Top of page
- Materials and methods
- Supporting Information
Experimental site description
The study was conducted on a sloping agricultural field and adjacent riparian zone located in the Thomas Brook Watershed of the Annapolis Valley, NS, Canada (N4504.886 W6445.318). The site has a cool temperate climate with humid soil moisture regimes and annual average (1971–2000) daily temperature and precipitation of 6.9 °C and 1211 mm, respectively. The lower part of the site (riparian zone) was a coarse-textured glacial fluvial Gleyed Humo-Ferric Podzol, while the upper part of the site (agricultural zone) was a finer-textured till Orthic Humo-Ferric Podzol. The 10-ha agricultural field was cropped to maize (Z. mays L.) while the riparian study zone was approximately 20 m wide and consisted of a mixed wood with herbaceous understorey (for details, see Leclair, 2010).
Four replicate transects were located in the study area, each of which had four sample positions based on landscape element within the agricultural zone and four sample positions within the riparian zone (eight positions for each transect, 32 positions in total) (Fig. 1). Slope positions were 1 (crest), 2 (shoulder), 3 (foot slope) and 4 (toe slope) in the agricultural zone and 5 (riparian edge), 6 (upper riparian), 7 (lower riparian) and 8 (brook edge) in the riparian zone. Management of the maize crop was conducted according to recommended industry practice. In brief, maize was seeded on 15 May 2007 with ammonium phosphate nitrate (N-P-K analysis of 30-10-0) banding at a rate of 200 kg ha−1, and ammonium nitrate (N-P-K analysis of 34-0-0) broadcast at a rate of 100 kg ha−1 on 3 July. Samples for this study were taken on three dates in 2007, representing different soil conditions: cool and moist soil conditions before second fertilizer application (7 June), after second fertilization and warmer soil temperature (10 July) and drier soil conditions (30 August). No irrigation was applied at the site.
Soil and gas sampling and analyses
On each sampling date, environmental variables (soil temperature at 10 cm depth taken using a Thermo Hygrometer Mannix Model CMM880; Cole-Parmer, Vernon Hills, IL) and water content from 0 to 15 cm soil depth (Hydrosense™ probe; Campbell Scientific, Edmonton, AB, Canada) were measured at each sample position.
Non-steady-state static vented chambers (Burton et al., 2008) were installed at each slope position in agricultural (between the maize rows) and riparian zones. Gas samples (20 mL) collected from each chamber at 0, 10, 20 and 30 min following deployment were stored in pre-evacuated 12-mL exetainers for transport to the laboratory. Gas samples were analysed by GC to determine N2O fluxes and respiration (CO2) from all the sample positions as described in Henderson et al. (2010).
Triplicate soil cores (0–15 cm) were sampled close to each flux chamber for inorganic N analyses and denitrification enzyme activity (DEA) assay (all dates), and total C and N, pH and soil texture (August only). Inorganic N species (NO3−-N+NO2−-N, NH4+-N) were extracted and analysed as described previously (Leclair, 2010). DEA was determined using the method described in Dandie et al. (2008). Total C and N were determined by combustion (LECO CNS-1000) (Bremner, 1996; Nelson & Sommers, 1996), soil pH was determined in a 1 : 1 soil : water suspension and soil texture was determined using the pipette method following organic matter removal.
Denitrification rates were determined using the acetylene blockage method as described by Paul & Zebarth (1997), from a separate set of soil cores (10 cm diameter × 15 cm) obtained from adjacent to the flux chambers at each sample position on all the sampling dates.
For nucleic acid extractions, six soil cores from each slope position were composited, homogenized and a subsample was placed into sterile 15-mL plastic tubes and immediately quick-frozen in dry ice in the field and then stored at −80 °C in the laboratory before further manipulation. Frozen soils were freeze-dried then DNA and RNA were coextracted from 1.5 g soil, divided equally into two 2-mL tubes as described in Henderson et al. (2010). The quality of DNA and RNA was assessed by agarose (1.0% w/v) gel electrophoresis. The lack of DNA in the RNA samples following DNase treatment was confirmed by performing a PCR reaction without reverse transcriptase. The RNA and DNA were quantified using the fluorescent dyes Ribogreen and Picogreen, respectively, as recommended by the manufacturer (Invitrogen, Burlington, ON, Canada).
Quantification of abundance and denitrification gene mRNA levels
Pseudomonas mandelii is a predominant culturable denitrifier from agricultural soil (Dandie et al., 2007a, b) and the abundance of nirSp gene-bearing denitrifiers (P. mandelii and related species) has been shown to be responsive to changes in soil conditions (Dandie et al., 2008; Miller et al., 2008); thus, it was included in this study. Gene copy numbers for nirS, nirK, nosZ and nirSp gene-bearing communities and transcript abundance for the nosZ gene-bearing communities were quantified via quantitative PCR (qPCR) and quantitative reverse transcriptase (qRT)-PCR using an Applied Biosystems (Streetsville, ON, Canada) ABI PRISM® 7000 thermal cycler and SYBR Green detection. PCR primers and conditions for amplification are described in detail in Table 1. Primers targeting nirS (Braker et al., 2000; Throbäck et al., 2004; Kandeler et al., 2006) and nirK (Henry et al., 2004) genes were used in qRT-PCR to quantify transcript abundance using a reverse transcription step of 30 min at 50 °C, followed by PCR amplification conditions and programme as described by the authors. No template controls were included in triplicate for all primer sets. Reactions not including reverse transcriptase were also included (nine random samples) in qRT-PCR runs to ensure that contaminating DNA was not present. Standard curves were constructed for the absolute quantification of denitrifier gene numbers and transcripts. Plasmids carrying cloned denitrification genes (Miller et al., 2008; Henderson et al., 2010) were prepared using standard methods, linearized using appropriate restriction enzymes and quantified using Picogreen (Invitrogen). Standard curves were generated using three replicates of 10-fold serial dilutions (from 101 to 106 copies) of plasmids containing the denitrifier gene sequences as a template and the cycling conditions described in Table 1. Soil DNA and RNA extracts were tested for the presence of coextracted inhibitory substances. A known quantity of the target gene or target mRNA was added to soil DNA or RNA extracts that were 10-fold serially diluted. The target gene was amplified using qPCR or qRT-PCR and the resulting gene copy or transcript numbers were compared with uninhibited samples. The template concentrations used were not found to be inhibitory to the qPCR or qRT-PCR reaction (data not shown).
|Primer name||Primer sequence (5′–3′)||Template||Primer concentration (nM)||Position||Target gene||Thermal profile||Reference|
|nirSCd3aF||AAC GYS AAG GAR ACS GG||1 μL 1 : 10 diluted DNA||500||916–935*||nirS||PCR – 95°C 10 min, 6 touchdown cycles [95°C 15 s, 63–58°C (−1°C per cycle) 30 s, 72°C 30 s, 80°C 30 s] then 40 cycles (95°C 15 s, 58°C 30 s, 72°C 30 s, 80°C 30 s)||Kandeler et al. (2006)|
|nirSR3cd||GAS TTC GGR TGS GTC TTS AYG AA||500||1322–1341*|
|nirKH1F||ATY GGC GGV AYG GCG A||1 μL 1 : 10 diluted DNA||1500||876–891b||nirK||PCR – 95°C 10 min, 6 touchdown cycles [95°C 15 s, 63-58°C (−1°C per cycle) 30 s, 72°C 30 s, 80°C 30 s] then 40 cycles (95°C 15 s, 58°C 30 s, 72°C 30 s, 80°C 30 s)||Henry et al. (2004)|
|nirKH1R||GCC TCG ATC AGR TTR TGG TT||2000||1021–1040†|
|nosZ1F||ATG TCG ATC ARC TGV KCR TTY TC||2 μL 1 : 10 diluted DNA||100||1184–1203‡||nosZ||PCR – 95°C 10 min, 6 touchdown cycles [95°C 15 s, 68-62°C (−1°C per cycle) 1 min, 81.5°C 30 s] then 40 cycles (95°C 15 s, 62°C 1 min, 81.5°C 30 s)||Henry et al. (2006)|
|nosZ1R||WCS YTG TTC MTC GAC AGC CAG||100||1415–1438‡|
|nirSsh2F||ACC GCC GCC AAC AAC TCC AAC A||5 μL 1 : 10 diluted DNA||300||226–247§||nirSp||PCR – 95°C 10 min, 40 cycles (95°C 15 s, 68°C 30 s, 72°C 30 s, 83°C 30 s)||Henderson et al. (2010)|
|nirSsh4R||CCG CCC TGG CCC TTG AGC||300||453–470§|
|nosZ1F||ATG TCG ATC ARC TGV KCR TTY TC||5 μL 1 : 10 diluted RNA||300||1184–1203‡||nosZ (RT-PCR)||RT – 50°C 30 min PCR – 95°C 10 min, 6 touchdown cycles [95°C 15 s, 68–62°C (−1°C per cycle) 1 min, 81.5°C 30 s] then 40 cycles (95°C 15 s, 62°C 1 min, 81.5°C 30 s)||Henry et al. (2006)|
|nosZ1R||WCS YTG TTC MTC GAC AGC CAG||400||1415–1438‡|
|nirScd3aF||AAC GYS AAG GAR ACS GG||300||916–935*||nirS||PCR – 94°C, 2 min, 40 cycles (94°C, 1 min, 58°C, 1 min, 72°C, 1 min), 72°C, 10 min.||Kandeler et al. (2006)|
|nirSR3cd||GAS TTC GGR TGS GTC TTS AYG AA||300||1322–1341*|
|Copper583F||TCA TGG TGC TGC CGC GKG ACG G||300||nirK||PCR – 94°C 5 min, 5 touchdown cycles [94°C 30 s, 72–67°C (−1°C/cycle) 1 min, 72°C 1 min] then 25 cycles (94°C 30 s, 67°C 1 min, 72°C 1 min), 72°C 7 min.||Wertz et al. (2006)|
|Copper909R||GAA CTT GCC GGT PGC CCA GAC||300|
Analysis of denitrifier community structures
Community structures of nirK- and nirS-bearing microorganisms in soil DNA extracts were characterized by denaturating gradient gel electrophoresis coupled with PCR (PCR-DGGE) analysis (see Table 1 for the PCR primers and amplification conditions). DGGE analysis of nirS and nirK PCR products used the D-code Universal Mutation Detection System (Bio-Rad) with a 6% polyacrylamide gel containing a gradient of 35–65% denaturant, 100% denaturing solution being defined as 7 M urea and 40% formamide. Gels were run for 4.5 h at 150 V in 1 × TAE buffer at 60 °C. The gels were stained with SYBR Green (Invitrogen) and then photographed.
Statistical analyses were conducted using the general linear model of systat (Systat Software Inc, Version 12, Chicago, IL). Non-normal data were log transformed. Soil pH, organic C and N and soil texture were analysed using the general linear model with slope position and transect (treated as replicates) as factors. Slope position means were compared using Tukey's honestly significant difference test. Comparisons between the agricultural and riparian land use zones were performed using single degree of freedom contrasts. Similar analyses were performed for other soil environmental properties and denitrifier community abundance and denitrification gene expression at each individual date. The treatment means and SEs presented in figures were calculated from untransformed data. Significance was accepted at P<0.05.
Multivariate statistical analyses
Similarities in the DGGE banding patterns among soil samples were represented by nonmetric multidimensional scaling (nMDS) (Kruskal & Wish, 1978). A data matrix consisting of the position and relative intensity of each DGGE band for all the samples was constructed using phoretix software (Phoretix International, Newcastle-Upon-Tyne, UK). For each sample, the ratios of the intensity of each band vs. the total band intensity were calculated. Rank similarity matrices (Bray–Curtis) were calculated from the data matrix using primer-e Ltd software version 6 (Plymouth, UK) and represented by nMDS. Distortions between the rank similarity matrices and the nMDS representations of similarities were assessed by calculating a stress value. Stress values from 0 to 0.2 indicate excellent to moderately good representation of the DGGE profile similarities by the nMDS. Significant differences in the community structure of denitrifiers between the agricultural and the riparian zones were evaluated using one-way analysis of similarity (anosim). anosim results in the computation of P-values providing the level of significance and R statistic values providing the degree of discrimination between the two zones and ranging from 0 to 1. R=0 if the similarities between and within replicates of the two zones are the same and R=1 if all the replicates of a zone are more similar to each other than to any replicates of the other zone.
Changes in bacterial denitrifier community structure (DNA banding patterns), denitrifier abundance and gene expression were related to the soil environmental properties and denitrifier activities using the BIO-ENV routine with Spearman rank correlation (Clarke & Ainsworth, 1993). All data points for individual dates and landscape positions were used in this analysis. Spearman rank correlation was used and 999 permutations of the data were used to test for the level of significance.
- Top of page
- Materials and methods
- Supporting Information
Denitrification, respiration and other soil physicochemical properties from three dates in the growing season presented in this study were excerpted from Leclair (2010). Soil texture, pH, total C and N were analysed by zone and slope position. Sand content was significantly higher in agricultural soil (54.8%) compared with riparian soil (46.0%), whereas silt content was higher in riparian soil (22.4%) compared with agricultural soil (15.7%). Clay content was similar between the two zones (Table 2). The average soil pH was slightly greater in the riparian zone (pH 5.50) compared with the agricultural zone (pH 5.15). Total C and N were significantly greater in the riparian zone by about twofold compared with the agricultural zone (Table 2). When the slope position was considered, soil texture did not change significantly and few changes in the soil C and N content were evident across slope positions (Table 2). Soil pH increased from 4.91 to 5.65 from slope positions 1 to 7 (Table 2). Because most soil properties did not change significantly as a function of slope position, we focused our attention on differences between land-use zones.
|Zone||Slope position||Soil texture||pH||Total N (g kg−1)||Organic C (g kg−1)|
|Sand (g kg−1)||Clay (g kg−1)||Silt (g kg−1)|
|Agricultural||1||54.3||30.1||15.7||4.91 a||0.087 a||1.83 a|
|2||56.2||29.7||14.1||5.05 ab||0.088 a||1.88 a|
|3||56.9||28.2||14.9||5.18 abc||0.072 ab||1.63 a|
|4||52.1||29.7||18.2||5.45 abc||0.080 ab||1.74 a|
|Riparian||5||48.6||33.7||17.7||5.55 bc||0.159 ab||2.76 abc|
|6||44.9||33.2||21.9||5.47 abc||0.159 ab||3.18 bc|
|7||43.8||30.4||25.7||5.65 c||0.230 b||4.83 c|
|8||46.8||29.1||24.1||5.38 abc||0.128 ab||2.68 ab|
Nitrate (NO3−) concentration, N2O flux, denitrification and water content varied significantly between agricultural and riparian zones (Table 3). Soil NO3− concentration was significantly greater in the agricultural zone (24.4–89.5 mg N kg−1 dry soil) compared with the riparian zone (1.9–4.3 mg N kg−1 dry soil; Table 3). Fluxes of N2O were 12–39 times greater in the agricultural zone compared with the riparian zone; however, the denitrification rate was greater in the riparian zone compared with the agricultural zone, with 1.6–5 times greater denitrification rate in June and July (Table 3). The water content was significantly higher in the riparian zone (0.27–0.44 g g−1 dry soil) compared with the agricultural zone (0.19–0.25 g g−1 dry soil). DEA was not significantly different between land-use zones in June and July, but was significantly greater in August in the riparian zone (90 mg N kg−1 dry soil day−1) compared with the agricultural zone (32 mg N kg−1 dry soil day−1). Although soil temperatures were generally cooler in the riparian zone (12.7–16.8 °C) compared with the agricultural zone (15.2–18.6 °C), only in June was there a significant difference in soil temperature between the two zones. Soil NH4+ concentrations and soil respiration rates were similar between the riparian and the agricultural zones (average of 3.5 mg N kg−1 dry soil and 13.7 g CO2 ha−1 day−1, respectively) (Table 3).
|NO3− (mg N kg−1dry soil)||24.4*||4.25*||89.5*||3.61*||41.4*||1.87*|
|NH4+ (mg N kg−1dry soil)||1.90||1.13||9.31||1.18||4.44||3.20|
|Respiration (kg CO2 ha−1 day−1||23.3||14.8||8.91||12.5||11.0||12.1|
|N2O flux (g N ha−1 day−1)||7.69*||0.65*||1.97*||0.16*||1.55*||0.04*|
|Denitrification (μg N kg−1 dry soil day−1)||40*||62*||5*||25*||15||26|
|DEA(mg N kg−1 soil day−1)||81||163||40||155||32*||90*|
|Soil temperature (°C)||15.2*||12.7*||18.3||14.5||18.6||16.8|
|Water content (g g−1 dry soil)||0.25*||0.44*||0.19*||0.27*||0.20*||0.31*|
Denitrifier abundance and denitrifier gene expression
Denitrifier gene (nirK, nirS, nirSp and nosZ) abundances were determined for each land-use zone (riparian and agricultural) at three time points in the growing season (Fig. 2). Average abundances for nirK ranged from 1.2 × 109 to 2.2 × 109 gene copies g−1 dry soil in the agricultural and riparian zones, respectively (Fig. 2a). Only in July was nirK abundance significantly higher (1.4-fold) in the riparian compared with the agricultural zone; however, at all the time points sampled, this trend was observed (Fig. 2a). The average abundance of nirS-gene-bearing denitrifiers was 2.1 × 106–1.7 × 106 gene copies g−1 dry soil in the agricultural and riparian zones, respectively (Fig. 2b). The abundance of nirS-gene-bearing denitrifiers was not significantly different between land-use zones for any of the individual three sampling dates (Fig. 2b). The average abundances of nirSp gene-bearing denitrifiers (targeting P. mandelii and closely related spp.) in the agricultural and riparian zones were 4.7 × 103–2.8 × 103 gene copies g−1 dry soil and were significantly higher (2.1-fold) in the agricultural compared with the riparian zone only in the June samples (Fig. 2c). In the agricultural and riparian zones, the abundance of nosZ gene ranged from 1.2 × 108 to 1.5 × 108 gene copies g−1 dry soil, respectively (Fig. 2d), and was not significantly different between land-use zones.
Primers targeting nirS- or nirK-bearing denitrifiers were not successful in qRT-PCR even after exhaustive optimization of the PCR reaction conditions and cycling parameters probably due to low detection levels or stability of transcripts and/or technical problems. Transcripts of the nosZ gene were quantified and compared between agricultural and riparian zones. Gene expression of nosZ was significantly greater by eightfold in the riparian compared with the agricultural zone in June and July, but did not differ between land-use zones in August (Fig. 2e). The lack of an effect of land-use zone on the gene expression of nosZ in August may reflect missing data points for several locations in the agricultural zone at this time (data not shown). Although mRNA could be extracted from all soil samples, reverse transcription and amplification was not obtained in all samples, despite multiple attempts and testing of alternative purification/optimization methods (data not shown).
There were no significant differences in nirK and nirSp abundance as influenced by slope position on any dates (data not shown). The nirS- and nosZ gene-bearing denitrifier abundance and the nosZ gene expression changed significantly along slope positions at specific dates, but only a few slope positions were significantly different and there were no logical trends in the data, with the exception of nirS abundance in June. There were significantly more nosZ-gene-bearing denitrifiers in the toe slope of the agricultural zone (position 4, 2.87 × 108 gene copies g−1 dry soil) than in the crest of the agricultural zone (position 1, 4.55 × 107 gene copies g−1 dry soil) in August. The nosZ gene expression was significantly higher in the lower riparian (position 7, 5.74 × 105 gene copies g−1 dry soil) than in the foot slope and toe slope of the agricultural zone (positions 3 and 4, with 1.44 and 2.33 × 105 gene copies g−1 dry soil, respectively) in June. The nosZ gene expression in the shoulder of the agricultural zone (position 2, 7.04 × 104 gene copies g−1 dry soil) was 23- and 27-fold lower compared with the riparian edge (position 5) and lower riparian (position 7) of the riparian zone in July. There were significantly more nirS-bearing denitrifiers in the toe slope of the agricultural zone (position 4), with 2.3 × 106 gene copies g−1 dry soil, compared with slope positions 1–7 (average of 4.81 × 105 gene copies g−1 dry soil), and there were significantly more nirS-gene-bearing denitrifiers in slope position 6 (upper riparian), with 7.58 × 105 gene copies g−1 dry soil, compared with 3.06 × 105 gene copies g−1 dry soil in position 8 (brook edge) of the riparian zone in June.
Denitrifier community structures
Changes in the community structure of nirS- and nirK-gene-bearing denitrifiers were evaluated by land-use zone and by slope position. Two-dimensional nMDS representations of community structure presented high stress distortion factors, which are probably due to high field variability (Fig. 3). Significant differences in the community composition between zones and levels of discrimination zones are indicated by R and P values (Fig. 3) The nirS community structure only differed between agricultural and riparian zones in June (R=0.22, P=0.002, Fig. 3a). Significant differences in the nirK community composition between the two zones were observed for all three sampling dates (R=0.25–0.31, P=0.001, Fig. 3b). Up to 15 bands for nirK- and up to 17 bands for nirS-gene-bearing denitrifiers were observed in the DGGE profiles. For the nirS gene DGGE profile, the relative abundances of bands 5, 16 and 8 differed significantly between agricultural and riparian zones. Bands 5 and 16 were present at greater relative abundances in the agricultural zone (i.e. 14.3% and 5.2%) compared with the riparian zone (i.e. 6.3% and 1.6%), and band 8 exhibited greater abundances (18.1%) in the riparian zone compared with the agricultural zone (11.5%) (Supporting Information, Fig. S1a). Similarly, for nirK gene DGGE profiles, the relative intensities of bands 1 and 2 were significantly higher in the agricultural zone (i.e. 8.4% and 12.6%) than in the riparian zone (i.e. 1.8% and 2.8%) and bands 11 and 12 showed greater relative abundances in the riparian zone (i.e 15.9% and 14.7%) compared with the agricultural zone (i.e. 7.9% and 9.1%) (Fig. S1b).
The nirS community structure differed among slope positions in June (R=0.17, P=0.02), but not in July and August, while the nirK community structure differed among slope positions on all three dates (R=0.16 and P=0.03 for June, R=0.24 and P=0.005 for July, R=0.17 and P=0.02 for August). Although some changes in nirS- and nirK-gene-bearing denitrifier community structure were noted, there were no gradual changes in denitrifier community structure when slope positions were considered (data not shown).
Relationships between denitrifier community structure, abundance, gene expression, denitrification and environmental parameters
BIO-ENV, using Spearman rank correlations, was used to relate denitrifier community structures, abundance and denitrification gene expression to denitrification and environmental characteristics of soil samples. The community structure of nirK gene-bearing denitrifiers was most strongly related to a single variable: soil pH (ρ=0.273, P=0.001, 999 permutations). nirS gene-bearing community structure, which did not change significantly between zones, was also not significantly correlated with any measured environmental variable when tested under these conditions.
No significant relationships were obtained between denitrifier abundances and denitrification or environmental conditions measured in this study, except for nirSp-gene-bearing denitrifier abundance, which was significantly related to the soil water content and soil NO3− concentrations (ρ=0.242, P=0.002, 999 permutations). No significant correlations were obtained between denitrifier community structure, abundance or denitrification gene expression and N2O fluxes, denitrification or DEA.
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This study evaluated changes in denitrifier abundance, nosZ gene expression, denitrifier community structure, denitrification and environmental properties along transects in a sloping landscape encompassing both agricultural and riparian in a cool temperate climate.
It was expected that changes in soil environmental properties with landscape position, independent of changes in land use, would influence denitrifier community abundance resulting in changes in denitrification and N2O emissions. Several studies on slope gradient reported important changes in soil properties including soil texture, organic C and pH (Pennock et al., 1992, 1994; Bronick & Lal, 2005). However, the experimental site of the current study showed limited differences in soil environmental properties as a function of slope position independent of the changes associated with differences in land use. A lack of response in soil environmental properties along a slope is not without precedence in Atlantic Canada (Zebarth et al., 2002). This study reported that soil environmental properties might not change if the slope in the landscape occurs on a site devoid of the soil A horizon probably due to extensive erosion in agricultural areas (Zebarth et al., 2002). Given that the experimental site did not show major changes in soil environmental properties among slope positions, it is not surprising that the abundance and diversity of the denitrifier communities targeted also did not respond or did not show a gradual pattern along slope positions.
Denitrification and environmental properties in agricultural and riparian zones
Environmental properties that differed between land-use zones included total organic C, total N, soil NO3− concentration, soil temperature and soil water content. Land-use zones varied in soil organic C concentration due to the depletion of organic C in the agricultural zone through soil disturbance (planting and tillage), soil erosion processes and the accumulation of C in the riparian zone through retention of organic matter in litter and limited disturbance to riparian soil. Total N in soils varied similarly with organic C concentrations. Crop fertilization resulted in higher soil NO3− concentrations in the agricultural zone compared with the unfertilized riparian zone. Soil temperature and soil water content are moderated by vegetation cover in the riparian zone, whereas the more exposed agricultural zone undergoes drier periods and warmer soil temperatures.
Denitrification rates were greater in the riparian compared with the agricultural zone. Studies have demonstrated that a high soil water content not only creates anoxic conditions that favour denitrification but also limits the diffusion of N2O away from the site of denitrification, thus resulting in N2 rather than N2O being the primary end product of denitrification (Knowles, 1982; Granli & Bøckman, 1994). Greater N2O fluxes were observed in the agricultural zone than the riparian zone, despite the greater denitrification rate in the riparian zone. This was attributed to the greater soil NO3− concentrations in the agricultural zone because high soil NO3− concentrations result in a greater proportion of gaseous emissions from denitrification as N2O emissions relative to N2, as NO3− is reduced preferentially over N2O (Betlach & Tiedje, 1981; Henderson et al., 2010). High rates of denitrification in the riparian zone have also been observed in other studies (Lowrance et al., 1984; Pinay et al., 1993; Hill, 1996). Similarly, Rich & Myrold (2004) observed greater potential of N2O formation in the riparian zone compared with the agricultural zone. It is also possible that nitrification was responsible for a portion of N2O emissions in the agricultural zone, given the form of fertilizer applied (ammonium nitrate) and the generally low water contents observed. Nitrification has been shown to contribute up to 81% of 15N-N2O at 60% water-filled-pore space (Bateman & Baggs, 2005); however, the contribution from nitrification to N2O emissions was not specifically assessed in this study.
Denitrifier abundances, nosZ gene expression and denitrifier community structures in agricultural and riparian zones
Denitrifier abundances in this study were monitored by qPCR. In comparison with other studies (i.e. Kandeler et al., 2006; Attard et al., 2010), nirK abundances were high (∼109 gene copies g−1 dry soil); however, similar high abundances have been reported previously by Chen et al. (2010) in a paddy soil using the same nirK qPCR primers (Henry et al., 2004). nirS gene abundances were much lower than for nirK, averaging ∼106 gene copies g−1 dry soil, which is within the range of nirS gene abundances reported previously (Yoshida et al., 2009). It is difficult to directly compare the gene abundances obtained in different studies, given the differences in DNA extraction efficiencies due to the soil type and/or the extraction method, different primer sets and differences in efficiency of the qPCR reaction (Smith & Osborn, 2009). It is notable in this study that the abundance of nirK genes was approximately 100–1000 times that of nirS genes and that this dominance of nirK-gene- over nirS-gene-bearing denitrifiers was consistent across the entire site sampled. Previous studies have observed a significant variation in ratios between nirS and nirK genes, with high nirK : nirS observed on few occasions (Bárta et al., 2010; Su et al., 2010). The reasons for the dominance of nirK-gene-bearing denitrifiers in these two ecosystems are not clear as there is limited information in the literature with regard to drivers of niche differentiation of nirK- or nirS-gene-bearing denitrifiers. A comprehensive study at the landscape scale determined pH to be the dominant explanatory variable for both nirK and nirS gene abundances, with total Cu being the second most important factor for nirK (Bru et al., 2010). Cu content was also identified by Enwall et al. (2010) as being significantly positively correlated with the quantity of nirK gene; however, the Cu content of soils in the region of our study was not measured and so we can only speculate whether this may have influenced the relative ratio of nirK : nirS we observed.
There were few changes in nirS-, nirSp-, nirK- and nosZ-gene-bearing denitrifier abundances between adjacent agricultural and riparian soil environments on any dates sampled, with the exception of nirK-bearing denitrifiers in July and nirSp-gene-bearing denitrifiers in June. Although a few studies measured the abundance of denitrifiers using most probable number (Hussey et al., 1985; McCarty et al., 2007) or C source utilization (Martin et al., 1999) in riparian soils, there is no information on the abundance of denitrifier communities in adjoining riparian and agricultural zones in the literature. The results of the current study provide evidence that the denitrifier community was both spatially and temporally stable in adjoining agriculture–riparian zones, despite differences in plant species and tillage practices between agriculture and riparian zones, which may have influenced the availability and quality of organic C, and other soil parameters. Further, despite the relative stability of the denitrifier communities, the rates of denitrification and N2O emissions were distinct between these two zones, indicating a disconnect between denitrifier abundances and associated process rates.
In contrast to nosZ-gene-bearing denitrifier abundance, there was significantly greater nosZ gene expression in the riparian than the agricultural zone. The nosZ gene expression is induced by anoxic conditions and the presence of NO (Arai et al., 2003; Vollack & Zumft, 2001) that is derived from the stepwise reduction of NO3− by the denitrification pathway in soils. Although soil NO3− concentrations were not high in the riparian zone, soil conditions were generally anoxic due to a high soil water content, which may be responsible for higher nosZ gene expression in this environment. Soil from the agricultural zone had high soil NO3− concentrations compared with the riparian zone; however, low nosZ gene expression in this zone suggests that induction of nosZ was limited by the soil oxic condition. Several studies have shown that anoxia is one of the major factors regulating denitrification through denitrification gene induction (Trevors, 1985; Tiedje et al., 1989; Saleh-Lakha et al., 2009). These results show that nosZ gene expression was more responsive to differences in environmental conditions than nosZ-gene-bearing denitrifier abundance.
Denitrifier abundances were relatively similar between riparian and agricultural zones; however, in contrast, particularly for nirK-gene-bearing denitrifiers, community structure was significantly different between land-use zones. Differences in the community structure between zones for the nirS-gene-bearing community were only apparent in June, indicating the relatively minimal response of this community to land-use differences. In contrast to the nirS community, the composition of the nirK community was different between the two land-use zones on all dates examined. Similarly, Rich & Myrold (2004) observed distinct nosZ-gene-bearing denitrifier community structures between a fertilized agricultural field planted to perennial ryegrass and a naturally vegetated riparian area. These results suggest that, in this study at least, not all denitrifier communities respond in the same manner to changes in the environment, and that certain groups of denitrifiers, i.e. the nirK-gene-bearing community, are more responsive to environmental differences between the agricultural and the riparian zones. The nirK-gene-bearing denitrifier community structure was also noted to be more responsive than nirS in different crop management systems (Enwall et al., 2010), and in land area representing a short-term plant succession (Smith & Ogram, 2008). The composition of the nirK-gene-bearing community in a potato field soil also changed significantly over a potato-growing season (Wertz et al., 2009).
Relationships between denitrifier abundance and community structure and environmental properties and denitrification
Attempts to relate denitrifier abundance and community structure to environmental properties or functional measures of denitrification or N2O emissions have been undertaken in a range of environments, with varying results (Rich & Myrold, 2004; Philippot et al., 2009; Wertz et al., 2009; Enwall et al., 2010). In this study, no significant relationships were obtained between nirS-, nirK- and nosZ gene-bearing denitrifier abundances or nosZ gene expression and environmental variables, despite measuring a range of soil conditions at the site. The exception was the nirSp community abundance (P. mandelii and closely related spp.), which had low, but significant positive correlations with both soil water content and soil NO3− concentration. Similarly, relationships between environmental properties and nirK- and nosZ-gene-bearing denitrifiers abundance were not observed in a grassland field (Philippot et al., 2009), with the exception of nirS abundance, which was related to several environmental factors including pH, NH4+ and NO3− concentrations and soil water content.
The community structure of denitrifiers bearing the nirS gene was not significantly correlated with any soil properties; however, the community structure of nirK-gene-bearing denitrifiers showed a correlation with soil pH. In a previous study in an organic and integrated cropping system, the opposite relationship was obtained, with nirS gene community structure being significantly correlated with pH, but nirK gene community structure was not (Enwall et al., 2010). Similarly, the narG-gene denitrifier community structure changed with soil pH in a legume field (Deiglmayr et al., 2004). Soil pH is often considered a master variable of bacterial community structure in soils and can explain a significant proportion of overall community structure; however, a large proportion of the variation in community structures remains unexplained (Lauber et al., 2009). Our results suggest that the nirK-gene-bearing denitrifier community was influenced by soil pH, even though the pH range in our soils was not large (0.74 pH units). Phylogenetic or physiological characterization of the dominant nirK-gene-bearing denitrifiers at low and high pH within the range observed in this study may provide some information on the mechanisms by which pH is influencing community structure in the two ecosystems, whether directly or indirectly, through the effect of pH on other soil properties. Further investigation or measurement of other soil parameters is required to gain an insight into the factors influencing denitrifier community structures in the two ecosystems.
There were no significant correlations between denitrifier abundance, nosZ gene expression or community structure and N2O fluxes, denitrification or DEA. Similarly, a number of studies have found denitrification rates, DEA and N2O emissions to be uncoupled from denitrifier abundance in potato field soil (Dandie et al., 2008), from denitrifier community structure in cultivated and uncultivated wetlands (Ma et al., 2008), in marsh sediment (Cao et al., 2008) and between riparian and agricultural zones (Rich & Myrold, 2004). In some instances, a relationship was observed between denitrifier abundance and N2O emissions or denitrification in estuary sediments (Dong et al., 2009) and in eight different crop systems with different management practices (Morales et al., 2010). Defining global parameters that link the abundance or the diversity of denitrifier communities and their gaseous emissions from soils may be difficult to achieve, given the phylogenetic diversity of microorganisms possessing this functional trait and the influence of soil physicochemical and other environmental factors on the processes involved. Alternatively, it may be the case that the abundance or the diversity of the denitrifier community may not be a rate-limiting factor in controlling the activity of denitrifying enzymes and as a result there may not be a consistent relationship between these parameters.
What is most remarkable about the observations made in this work is the relative stability of denitrifier community abundance in land-use zones showing different plant species, soil conditions, DEA and denitrification rates. Nonetheless, the denitrifier community structure of some of the denitrifier communities did change. This study demonstrated that nirK community structure was more responsive than nirS community structure to change between agricultural and riparian zones. There was a clear relationship between nirK community structure and pH, and between nirSp, abundance and soil NO3− concentration and soil water content. The results also revealed that changes between denitrifier abundance and community structure were uncoupled. This suggests that different processes define the niche that denitrifiers occupy and the factors controlling their abundance, relative to the diversity of denitrifiers that occupy that niche. The expression of the nosZ gene was influenced by higher soil water content and anoxia associated with the continuous vegetation cover of the riparian zone, which helps to explain the higher denitrification observed in this zone.
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Jan Zeng and Drucie Janes are acknowledged for excellent help in field sampling and for conducting laboratory molecular, soil and gas analyses. We thank the anonymous reviewers for improving the manuscript. Funding for this work was provided by the GAPS program of Agriculture and Agri-Food Canada and an NSERC (Canada) Strategic Team Grant. Research by J.T.T. is also funded by an NSERC (Canada) Discovery grant.
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Fig. S1. Representative examples of DGGE profiles of (a) nirS genes for the samples of June (transects 1 and 4) and (b) nirK genes for the samples of July (transects 1 and 2).
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