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DIVISION S-10-WETLAND SOILS

Influence of Nitrate and Phosphorus Loading on Denitrifying Enzyme Activity in Everglades Wetland Soils

^a Univ. of Florida, Wetland Biogeochemistry Lab., 106 Newell Hall, P.O. Box 110510, Gainesville, FL 32611 USA

krr{at}gnv.ifas.ufl.edu

	ABSTRACT

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
There has been recent concern about the impact of increased nutrient loading on the northern Everglades ecosystem. We investigated the spatial and temporal distribution of denitrifying enzyme activity (DEA) along a P-enrichment gradient in the Water Conservation Area 2A (WCA-2A) and determined the effects of added P and NO^-₃ on DEA. The DEA in soil and detritus layers was measured under anaerobic conditions four times during 2 yr, using the acetylene blockage technique. The DEA ranged from 0.004 to 7.75 mg N₂O–N kg^-1 h^-1. Highest rates of DEA were found in the detritus and surface (0–10 cm) soils, and rates decreased exponentially with increasing distance from the surface-water inflow point, where nutrients are loaded to the wetland. Nitrate was found to be limiting, while the addition of P had no effect on the distribution of DEA in these soils. There was a seasonal effect on DEA, with higher activity observed during the summer when temperatures and hydraulic and nutrient loading were highest. Soils from outside the impacted zone demonstrated denitrifying potentials, within 10 h when spiked with inflow concentrations of NO^-₃, similar to DEA of soils from within the impacted zone. This suggests that soils from outside the impacted zone can increase denitrification rates when exposed to higher NO^-₃ concentrations in a relatively short time. Agricultural drainage water discharge, and consequent NO^-₃ loading, has created a zone of elevated DEA proximal to the S-10C surface-water inflow point in WCA-2A.

Abbreviations: ANOVA, analysis of variance • DEA, denitrifying enzyme activity • SFWMD, South Florida Water Management District • WCA-2A, Water Conservation Area 2A

	INTRODUCTION

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
NITRATE REDUCTION is the major N removal mechanism in wetlands. Among NO^-₃ reduction processes, denitrification is the dominant NO^-₃ removal process. Denitrification is a microbially mediated process whereby facultative anaerobic bacteria use NO^-₃ (or NO^-₂) in the absence of O₂ as the terminal electron acceptor during the oxidation of organic C (microbial respiration), resulting in the production of gaseous end products, N₂O and N₂. The denitrification enzyme assay is used as a means to eliminate all other regulating factors of denitrification in order to quantify the amount of active denitrifying enzymes present in soil (Smith and Tiedje, 1979; Smith and Parsons, 1985; Groffman, 1987; Schipper et al., 1993). The enzyme assay is the short-term (2 h) rate of N₂O production and is indicative of the size and activity of the denitrifying enzyme pool present in soil. The assay reflects the immediate biological effect of changes in redox conditions attributed to changes in soil O₂ levels (Martin et al., 1988).

Several studies of DEA have focused on upland soils (Smith and Tiedje, 1979; Groffman, 1987; Parsons et al., 1991), with the goal of more recent studies focused on correlating denitrification potential at the ecosystem scale to easily measurable soil parameters. These studies have reported rates of N₂O production in upland soils ranging from 0.006 to 7.14 mg N kg^-1 h^-1. The results of such research could be used to quantify the contribution of soils to global atmospheric N₂O levels.

Several problems exist with the use of DEA soil measurements in extrapolating to landscape scale denitrification rates in upland soils. The microorganisms responsible for the production of enzymes are facultative anaerobes, which possess separate enzyme systems capable of using either O₂ or NO^-₃ as terminal electron acceptors. The NO^-₃ reducing enzyme systems are primarily inactive in the presence of O₂ and active in enzyme production only during ephemeral anoxic events (e.g., rainfall events; Sexstone et al., 1985; Burton and Beauchamp, 1985). Further, the presence of O₂ appears to repress or deactivate enzymes already present in soil (Martin et al., 1988). Secondly, the distribution of organic matter in upland systems is patchy, which presents additional problems in assessing the spatial distribution of DEA. Hotspots of organic matter provide both simple C compounds for the maintenance of large, microbial populations and anoxic microenvironments that lead to NO^-₃ consumption as the terminal electron acceptor (Parkin, 1987; Christensen et al., 1990a, 1990b). The patchy distribution of active enzyme sites in the landscape leads to logarithmic frequency distributions of enzyme activity measured in the field (Parkin, 1987). Finally, NO^-₃ is rarely the limiting factor for denitrification in upland ecosystems, as evidenced by the widespread problem of groundwater contamination of NO^-₃, and consequently is a poor indicator of denitrification potential. These factors in combination confound efforts to consistently correlate easily measured soil parameters (total C, water content, NO^-₃ concentration, and microbial biomass) with DEA to produce meaningful estimates of denitrification in the landscape. (Parsons et al., 1991; Velthof et al., 1996).

There exist several important differences between mineral, upland soils and organic-rich, wetland soils that permit the reliable use of DEA on a landscape scale in order to investigate the source and effect of NO^-₃ loading in wetlands. Wetland soils are often saturated most of the year, thereby reducing diffusion of O₂, resulting in all but the smallest (1–4 mm) surface layer of soil remaining anaerobic (DeBusk and Reddy, 1998). Christensen et al. (1990b) found that the frequency distribution of denitrification rates was less skewed in upland soils after the onset of flooded conditions. This paucity of O₂ prevents the repression of denitrifying enzymes present. The relatively high organic matter content of wetland soils provides ample substrate for heterotrophic microbial activity and any O₂ that contacts the soil is quickly used. These conditions suggest that NO^-₃ supply is the limiting factor for denitrification in wetlands (Cooper, 1990) and that the presence of NO^-₃ will therefore control the size and activity of the denitrifier microbial populations. Schipper et al. (1993) found that up to 77% of the variability of in situ denitrification in an organic riparian wetland soil could be explained by NO^-₃ concentration and DEA. They also noted that organic soils, which comprised only 12% of the total catchment, were responsible for 56 to 100% of total NO^-₃ consumption. Therefore, it has been suggested that an increase in DEA in flooded soils can be in response to increased NO^-₃ loading and that DEA might be a valuable indicator for determining the areal extent of impact of NO^-₃ additions in wetlands.

	Study Area

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
The Florida Everglades are currently affected by nutrient loading from urban and agricultural surface runoff. Most notably, this impact is seen in the Water Conservation Areas, one of the major hydrologic units of the Everglades (DeBusk et al., 1994). Water Conservation Area 2A has been receiving nutrient-laden (N and P) drainage waters for the past 40 yr. Peat and nutrient (organic C, N, and P) accretion rates have increased in areas receiving surface drainage water (Koch and Reddy, 1992; Craft and Richardson 1993). Most notably, the effect of anthropogenic nutrient loading is documented in the spatial distribution of surface soil total P. Total P concentration grades from a high of 1600 mg kg^-1 at the surface-water inflow points to a background concentration of 400 mg kg^-1 in unimpacted areas located in the interior of the marsh (Koch and Reddy, 1992; Reddy et al., 1993; DeBusk et al., 1994). A gradient in NO^-₃ and soluble P concentrations in the water column and periphyton tissue has also been documented along the same transect in WCA-2A (McCormick and O'Dell, 1996). Historically an oligotrophic, P-limited sawgrass (Cladium jamaicense Crantz) marsh, the vegetation began a shift towards a dominant cattail (Typha domingensis Pers.) vegetative community proximal to all surface-water inflow points (Davis, 1991; Craft and Richardson, 1997).

The objectives of this study were to determine (i) spatial and temporal distributions of DEA for wetland soils of WCA-2A, (ii) effects of added P and NO₃–N on DEA, and (iii) the relationship between soil properties and DEA.

	Materials and methods

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
Experimental Design
Ten stations were located along a 10-km transect originating from the S-10C inflow water control structure in WCA-2A (Fig. 1) spanned the marsh from a primary water control inflow structure (S-10C) southward across the dominant cattail vegetation and terminated 10 km into the natural (unimpacted) marsh. The natural marsh was characterized by stunted stands of sawgrass separated by shallow sloughs dominated by floating and attached cyanobacterial mats. Sampling stations were located at distances of 0.2, 0.3, 1.4, 2.3, 3.3, 4.2, 5.1, 7.0, 8.4, and 10.1 km from the S-10C water control structure. Water depths varied seasonally from <2 cm to 2 m along the transect length in 1995 and 1996 (South Florida Water Management District [SFWMD], 1996, unpublished data).

View larger version (19K): [in this window] [in a new window]   Fig. 1 Station locations along a soil P gradient in WCA-2A (south of the S-10C water control structure) used in this study

The SFWMD established 21 circular tanks (mesocosms) enclosing 1.8 m² and three open control plots in an unimpacted sawgrass–periphyton slough in order to isolate the effects of added P on soil characteristics and periphyton (McCormick and O'Dell, 1996). The mesocosm site was located 11 km southwest of the S-10C inflow water control structure (Fig. 1). The mesocosms were installed entirely within a shallow slough that contained no established stands of sawgrass within the study site. The soil surface was dominated by floating and benthic cyanobacterial (periphyton) mats, purple bladderwort (Utricularia purpurea Walt.), and water lily (Nymphaea odorata Ait.) (McCormick and O'Dell, 1996). Three replicate mesocosms were selected at random and spiked once a week with various amounts of NaH₂PO₄ mixed with slough water to achieve annual loading of 0, 0.4, 0.8, 1.6, 3.2, 6.4, and 12.8 g P m^-2 yr^-1. The tanks were closed from exchange with the surrounding water by sliding a plastic collar over the holes for 24 h after spiking. The tanks were subsequently opened to permit exchange with the surrounding slough during the no-dose periods. Prior to our soil sampling, these systems had been dosed weekly at respective levels for 17 mo.

Soil Sampling and Characterization
A minimum of four soil cores were collected within 5 m of each station along the transect by driving a 10-cm-diameter aluminum irrigation pipe into the soil. A probe was inserted into each core to verify that negligible (<5%) compaction had occurred during coring. Cores were sealed, removed from the ground, immediately extruded, and separated into separate soil intervals (0–10 and 10–30 cm) in the field. Each interval was well mixed to yield a representative and homogenous sample from each depth at each station. The August 1995 and February 1996 samples were bagged and immediately transported on ice to the laboratory in Gainesville, FL. Samples were transferred into 2-L polyethylene containers within 24 h of collection and stored refrigerated at 4°C until analysis. Soil samples collected in August 1996 and March 1997 were immediately transported to a field laboratory location and incubated within 3 h of collection. Detrital surficial litter material was collected during the last two sampling events for use in field incubations. Detritus consisted of recognizable, loosely associated cattail or sawgrass plant material lying on the surface of the underlying more compact, brown peat soil. The detrital layer varied in thickness from <1 cm in sawgrass areas to >25 cm at the cattail stations closest to the inflow. The remaining soil samples not used in field incubations were sealed in plastic bags and kept on ice until return to the laboratory, where the samples were transferred into polyethylene containers and refrigerated at 4°C until subsequent characterization.

Soils were collected from experimental mesocosms on 21 Nov. 1996 by driving a 10-cm-diameter polyethylene tube into the soil. The periphyton-floc layer was poured off into separate sampling containers. The top soil interval (0–3 cm) was then extruded, stored in plastic bags, and placed on ice until returning to the laboratory, where samples were stored refrigerated at 4°C until subsequent characterization.

Bulk density was calculated for the soil intervals on a dry weight basis. Total C and N contents of detritus and soils were determined on dried, ground samples using a Carlo-Erba NA-1500 CNS Analyzer (Haak-Buchler Instruments, Saddlebrook, NJ). Total P analysis was performed on subsamples prepared by nitric-perchloric acid digestion (Kuo, 1996) and analyzed using an automated ascorbic acid method (Method 365.4; USEPA, 1983).

Microbial biomass C was determined using the fumigation-extraction technique of Vance et al. (1987) for the February and August 1996 and March 1997 sampling times. Six replicate 5-g samples were placed into 25-mL centrifuge tubes for each soil interval and all 10 sampling sites. One-half milliliter of chloroform was added to three replicate tubes and placed in a vacuum desiccator with a beaker containing 300 mL of chloroform and several boiling chips. After 24 h, the headspace was alternatively evacuated and filled with room air nine times to remove chloroform still present in the soil or beaker. Samples were removed and both the controls (not exposed to chloroform) and chloroform-treated soils were immediately extracted with 20 mL of 0.5 M K₂SO₄, agitated for 30 min on a longitudinal shaker, and vacuum filtered through no. 42 Whatman filter paper. The supernatant was collected and refrigerated at 4°C until analyzed on a Dohrman total organic C analyzer (Rosemount Analytical, Santa Clara, CA). Microbial biomass was determined by subtracting the extractable total organic C in the triplicate controls from the triplicate chloroform-treated samples. An extraction efficiency (k_EC) factor of 0.37 was applied, using a previous calibration by Sparling et al. (1990) for organic soils.

Denitrifying Enzyme Activity
Laboratory DEA incubations were performed on soils collected along the transect in August 1995 and February 1996 and on soils collected from the mesocosms in November 1996. Approximately 10 g of field-moist soil from each site and depth were placed in triplicate 110-mL glass serum bottles and sealed with rubber septa and aluminum crimp caps. The headspace was evacuated to -85 kPa and replaced with O₂-free N₂ gas to achieve anaerobic conditions. Five milliliters of N₂-purged distilled, deionized water were added to each bottle to create a soil slurry. Approximately 15% of the headspace N₂ was replaced with acetylene gas (C₂H₂) (Balderston et al., 1976; Yoshinari and Knowles, 1976). Bottles were shaken on a longitudinal shaker for 1 h to evenly distribute the C₂H₂ throughout the soil slurry. Eight milliliters of solution containing 56 mg NO₃–N L^-1, 288 mg C₆H₁₂O₆–C L^-1, and 2 mg L^-1 chloramphenicol (an enzyme production inhibitor) were added to each bottle, creating a slight overpressure (Smith and Tiedje, 1979). Samples were incubated in the dark at 25°C and continually agitated on a longitudinal shaker. Headspace gas was sampled every 30 min for 2 h. Nitrous oxide production was adjusted for N₂O dissolved in the aqueous phase using a Bunsen absorption coefficient of 0.544 for N₂O (Tiedje, 1982). The denitrification rate was calculated by determining the slope of the linear curve produced when cumulative N₂O evolution was plotted vs. time.

Field DEA incubations were performed on freshly collected soils (during August 1996 and March 1997) within 3 h of sampling. Incubations followed the same procedures as those performed in the laboratory, with the following modifications because of field constraints. Bottles were evacuated by pulling a 60-cm³ syringe three times to evacuate the headspace and incubated submerged in site water at ambient temperatures (29–31°C) without shaking. Also, headspace gas was sampled at the terminus of the 2-h incubation, placed in evacuated 3.5-mL serum bottles sealed with butyl rubber stoppers and aluminum crimp caps, and transported to the lab for subsequent gas analysis within 48 h.

Denitrifying Potential
Surface soils (0–10 cm) from Station 1 (located 0.2 km from the water control structure) and Station 10 (located 10.1 km from the water control structure) were subjected to inflow water concentrations of NO^-₃ (1 mg L^-1 NO₃–N) under a 15% C₂H₂ (v/v) headspace for 24 h without addition of an enzyme inhibitor or exogenous C in order to determine the denitrifying potential of soils from inside and outside the impacted region in response to NO^-₃ loading. Samples were continuously shaken at 25°C in the dark to negate diffusion limitations. Headspace gas was sampled periodically until the N₂O production curve flattened out, indicating the complete consumption of added NO^-₃. The potential denitrification rate at this inflow floodwater NO^-₃ concentration was calculated from the steepest part of the N₂O production curve.

Nutrient Addition Study
Surface soil (0–10 cm) from Station 10 (10.1 km from the inflow) was collected to determine the effect of added KNO₃ and NaH₂PO₄ on DEA. The soil was homogenized by mechanical mixing after removing live roots. Approximately 50 g of field-moist soil was added to each 120-mL media bottle equipped with a butyl rubber septum embedded in the cap. To each bottle, 40 mL of distilled, deionized water was added and mixed well with the soil. The following treatments were evaluated:

1. Control (no additions)
   
2. Added NO₃–N (1, 10, 50, 100 mg L^-1 porewater concentration)
   
3. Added PO₄–P (0.1, 1, 5, 10 mg L^-1 porewater concentration)
   
4. Added NO₃–N + PO₄–P (matching rates as shown in Treatments 2 and 3, with the lowest rate of Treatment 2 added in conjunction with the lowest rate in Treatment 3 and so forth)
   

Each treatment was performed in triplicate. Bottles were capped and purged with O₂-free N₂ gas to induce anaerobic conditions. Samples were incubated in the dark at 30°C for 10 d and were shaken by hand for 30 s d^-1. Bottles were then respiked with the same concentration of the respective solutions and incubated for an additional 10 d. A triplicate set of soil controls was spiked with distilled, deionized water and was included in the incubation. At the terminus of 20 d of incubation, sample bottles were opened and 20 mL of slurry was collected from each bottle and placed in a serum bottle under a N₂ headspace containing 15% C₂H₂. Potential denitrification rates were determined without NO^-₃ additions for 48 h for all samples. An additional 20 mL of soil slurry was subjected to the denitrification enzyme assay procedure to investigate the effects of nutrient additions on soil DEA values.

Gas Analysis
Gas samples were analyzed for N₂O on a Shimadzu (GC-14A, Shimadzu Scientific, Kyoto, Japan) gas chromatograph equipped with a 3.7 x 10⁸ (10 mCi) ⁶³Ni electron capture detector (300°C ). An 1.8 m by 2 mm i.d. stainless steel column packed with Poropak Q (0.177–0.149 mm; 80–100 mesh) was used (Supelco, Bellefonte, PA). The carrier gas (5% CH₄ in Ar; v/v) flow rate was 30 mL min^-1 maintained at 30°C. Working standards consisted of N₂O in a framework of He gas (Scott Specialty Gas, Plumsteadville, PA).

Data Analysis
Data were fitted to an analysis of variance (ANOVA) model to investigate significant differences (P < 0.05) in DEA among stations, soil depths, seasons, and nutrient addition levels. A paired student's t test was used to detect significant differences (P < 0.05) between seasonal distributions of DEA along the transect length. The DEA also was statistically correlated with soil characterization data to determine which soil parameters were the best predictor(s) of enzyme activity. Fisher's Least Significant Difference (LSD) test was used to make comparisons among treatment levels for the nutrient addition study, using the StatGraphics software program (Manugistics, Rockville, MD).

	Results and discussion

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
Soil Characterization
Average bulk densities of organic soils collected along the transect for all samplings were 0.064 and 0.089 g cm^-3 for the 0- to 10-cm and 10- to 30-cm soil depths, respectively (Table 1) . Bulk densities were not determined for detritus samples. Total C and N did not vary significantly along the transect, but total P was significantly negatively correlated (P < 0.01) with distance from the inflow for detritus, 0- to 10-, and 10- to 30-cm soil depths (r = -0.882, -0.971, and -0.833, respectively). Results of a one-way ANOVA showed that total P was significantly higher (P < 0.05) in both detritus and 0- to 10-cm soil when compared with the underlying 10- to 30-cm soil, while there was no significant difference in total P content of detritus and the 0- to 10-cm soil interval. Microbial biomass C was significantly negatively correlated with distance (r = -0.68; Table 2) . A positive correlation was observed for microbial biomass C vs. total P (r = 0.65) for the detrital layer. The microbial biomass C decreased with depth along the transect (P < 0.01; r = 0.69). Microbial biomass C averaged 13.3, 4.8, and 1.6 g kg^-1 for the detritus, 0- to 10-, and 10- to 30-cm soil depths, respectively.

View this table: [in this window] [in a new window]   Table 1 Select physiochemical properties of detritus and soils collected from along the study transect in WCA-2A.

View this table: [in this window] [in a new window]   Table 2 Product-moment correlation coefficients for selected parameters for soil samples collected from along the study transect in WCA-2A.

View this table: [in this window] [in a new window]   Table 3 Select physiochemical properties of the 0- to 3-cm soil interval from the mesocosm experiment in WCA-2A.

View larger version (22K): [in this window] [in a new window]   Fig. 2 Denitrifying enzyme activity vs. distance for the 0- to 10- and 10- to 30-cm soil depths for the August 1995 and February 1996 samplings from the transect in WCA-2A. Plotted are mean values and one standard error

View larger version (21K): [in this window] [in a new window]   Fig. 3 Denitrifying enzyme activity vs. distance for the 0- to 10- and 10- to 30-cm soil depths for the August 1996 and March 1997 sampling from the transect in WCA-2A. Plotted are mean values and one standard error

Denitrifying enzyme activity of detritus and surface soils within 2 km of the inflow in WCA-2A are among the highest rates reported in the literature (Table 5) and are attributed to the stimulation of denitrifying bacteria by the inflow concentrations (1 mg N L^-1) of NO^-₃ in agricultural drainage waters. Lower levels of DEA at stations furthest from the inflow are probably related to nitrification processes in the overlying water column, producing NO^-₃ from NH⁺₄ which then diffuses into the soil under a concentration gradient where it can be used by denitrifiers (Reddy and Patrick, 1984).

The results of a paired student's t test suggest higher DEA for the summer or wet season (August 1995) 0- to 10-cm soil data than for the winter or dry period (February 1996) at P < 0.05. A similar trend was evident in the 10- to 30-cm soil depth at P < 0.05, with higher DEA in August 1995 than in February 1996. Highest DEA was found closest to the surface-water inflow point during the summer months, further supporting the hypothesis that increased soil DEA is stimulated by NO^-₃ loading. There was no significant difference (ANOVA) between sampling times for the 10- to 30-cm soil interval, suggesting NO^-₃ supply to the subsurface soils is supplied primarily through nitrification processes at the soil–root interface as opposed to diffusion of NO^-₃ down from the water column.

The seasonal differences in DEA appear to be related to the surface-water management schedule. The SFWMD maintains increased surface-water flow into the WCA-2A during the wet summer periods. Both studies in summer (August 1995, 1996) yielded higher N₂O production rates at stations closest to the inflow as compared with the same sites during the winter (February 1996, March 1997). As the majority of NO^-₃ loading occurs in the summer (wet season) months, there appears to be a concomitant increase in denitrifying enzyme activity.

Impact of Nitrate Loading on Denitrifying Enzyme Activity
The areal extent of relatively high DEA values appears to be confined within 2 to 4 km of the inflow point, with little apparent change along the transect after that point. The average daily NO^-₃ removal rate of soil layers within 2 km of the inflow point (using DEA values) was calculated to be 2.22 x 10⁶ g N d^-1 for 1995. This estimate was determined calculating the average DEA for the entire soil volume contained radially within 2 km of the inflow point. The daily loading rate of NO^-₃ to WCA-2A through the S-10C water control structure as determined by the SFWMD in 1995 was 2.50 x 10 ⁶ g N d^-1 (SFWMD, 1995, unpublished data). The apparent agreement between these two estimates suggests all of the NO^-₃ loaded to the system could potentially be removed by denitrifying enzymes present in the soil.

Potential denitrification rates are generally an overestimation of field or in situ rates in uplands, as diffusion constraints are present in the field. However, Shipper et al. (1993) found that DEA and field rates of ambient denitrification were similar in an organic riparian wetland. Regardless, hydraulic loading to WCA-2A is sporadic or discontinuous, so it is likely that a plume of inflow water could extend beyond the defined impacted region during high surface-water flow events. Therefore, we investigated the effect of inflow concentrations of NO^-₃ on denitrifying potential of soil samples located away from the region of elevated DEA and compared those rates with DEA of soil within the impacted area.

Soil samples were subjected to inflow concentrations of NO^-₃ (1 mg N L^-1) from within the impacted region reduced the added NO^-₃ to N₂O gas within 4.5 h of the start of the incubation (Fig. 4) . The denitrification rate was linear and approximated the short-term DEA rate. The unimpacted soil demonstrated low initial denitrification rate similar to that for the 2-h enzyme assay. However, stimulated by the added NO^-₃ across 10 h, denitrifying enzyme activity increased the N₂O production rate to the same order of magnitude as the DEA rates from the impacted site. This result demonstrated that soils from outside the impacted area are capable of quickly responding to increased NO^-₃ loading by increasing biological denitrification rates. Cooper (1990) reported high DEA along the upslope edge receiving NO^-₃ inputs to an organic riparian wetland, yet low or undetectable levels of DEA further downslope. The author concluded that the organic soils downslope were capable of higher denitrification, but were simply NO^-₃ limited. A similar conclusion was reached by Schipper et al. (1993).

View larger version (15K): [in this window] [in a new window]   Fig. 4 Cumulative N2O production curves for the 0- to 10-cm soil depth located 0.2 km (impacted) and 10.1 km (unimpacted) from the water control inflow structure in WCA-2A. Plotted are means (n = 4) and one standard error. Samples were collected in March 1997

View larger version (14K): [in this window] [in a new window]   Fig. 5 Denitrifying enzyme activity vs. total P for detritus and the 0- to 10- and 10- to 30-cm soil depths along the transect in WCA-2A

Surface soils (0–3 cm) in experimental mesocosms had been loaded with variable rates of P by the SFWMD (McCormick and O'Dell, 1996). Distribution of DEA values in the mesocosm study showed no significant correlation with total soil P, although total P was significantly (P < 0.01) correlated with P-loading rate (Table 4). However, the microbial biomass C was also significantly higher in soils with higher total P contents (r = 0.50). This suggests a P limitation to the microbial pool in natural Everglades peat soils. The fact that microbial biomass C and DEA are not significantly correlated could be due to the existence of a wide variety of functional microbial groups present in the soil (Drake et al., 1996). Denitrifiers make up a small percentage of the total microbial biomass where very little NO^-₃ or O₂ is available to sustain microbial respiration. Duncan and Groffman (1994) also found no significant correlation between microbial biomass C and DEA for a natural and treatment wetland.

These results led to the conclusion that the strong correlation of DEA with total P along the transect was simply an incidental relationship rather than causal. Nitrate N appeared to be the limiting factor as both N and P are loaded to WCA-2A in the drainage water. Phosphorus did not appear to be limiting to the denitrifier populations in the soil at the background level of NO^-₃ loading (supplied through nitrification). If NO^-₃ were loaded at sufficiently high concentrations so as not to be limiting, perhaps a P limitation might then be expressed.

Nutrient Addition Study
Potential denitrification rates, determined on surface soil samples spiked and incubated with various levels of NO₃–N, PO₄–P, and NO₃–N + PO₄–P, demonstrated significant differences across the range of N and N + P additions, but no significant differences for all levels of P. In addition, the two lowest levels of N and N + P additions also demonstrated no significant difference in denitrification rate compared with the control (no addition) or P-only additions. Significant differences were only noted at the two highest levels of N (Table 6) . This suggests that denitrification in Everglades soils is controlled by NO^-₃ input, not P. Interestingly, there was also significantly higher denitrification for soils spiked with the two highest levels of N and P, compared with samples spiked solely with the same levels of N only. This suggests that as N becomes nonlimiting, P can become the limiting nutrient for denitrification. There were no significant differences between samples that had undergone N + P and N additions at the two lowest levels, suggesting NO^-₃ limitation to denitrification.

Conclusions
The results of this study suggest that present NO^-₃ loading rates to WCA-2A wetland have had an effect on the spatial and temporal distribution of soil DEA. Data suggest that the majority of the exogenous NO^-₃ is intercepted by this microbial pool and lost from the wetland as gaseous N₂ and N₂O. The majority of NO^-₃ reduction occurs in the detrital and surface (0–10 cm) soil layer. The distribution of DEA in surface soil decreased along the transect as a first-order decay function. During high-loading seasons (summer), DEA distributions fit this exponential equation; however, during periods of low NO^-₃ loading (winter), DEA distribution did not fit either a zero- or first-order model. This suggests that soil denitrifying enzymes are produced in response to increased NO^-₃ loading.

Further investigations suggest that increased hydraulic loading into WCA-2A, thereby increasing NO^-₃ loading, would stimulate a concomitant increase in the activity of denitrifying populations in soils and detritus. This increase in enzymes activity or production would aid in the removal of additional NO^-₃ associated with increased loading. Phosphorus loading appeared to have minimal effect on the level of soil DEA, leading to the conclusion that the strong correlation between DEA and total P in the WCA-2A wetland is coincidental and not causal. A P limitation on denitrifying potential and DEA was demonstrated at highest levels of NO^-₃ additions, significantly higher than N concentrations of either the impacted or unimpacted soils. This suggests in situ rates of NO^-₃ reduction of Everglades soils and detritus are NO^-₃ limited in WCA-2A.

This study suggests WCA-2A could potentially receive a far greater NO^-₃ load without reaching a saturation limit on the potential denitrification capacity of soils within this 44684-ha wetland. It has been demonstrated that the denitrifying enzyme assay can be used to discern areas of increased NO^-₃ loading in flooded soils. This application of DEA might be used in other aquatic systems (e.g., lakes and streams) to identify soils and sediments that are intercepting plumes of NO^-₃ carried by surface-water flow or by groundwater; however, a calibration to determine baseline-level DEA characteristics may be needed for each system.

	ACKNOWLEDGMENTS

 
This study was supported in part by the South Florida Water Management District, West Palm Beach, FL. The authors would like to acknowledge the assistance of Drs. Paul McCormick and Sue Newman during the course of field investigations, especially for allowing use of their experimental mesocosms. In addition, the authors acknowledge the assistance of Dr. Bill DeBusk, Matt Fisher, and Yu Wang during the course of this investigation. We also thank Dr. Brian McNeal, Dr. Don Graetz, and several anonymous reviewers for their constructive critical comments, which greatly improved the quality of the manuscript.

	NOTES

TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 
Florida Agricultural Experiment Station Journal Series no. R-06680.

Received for publication December 17, 1998.

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TOP NOTES ABSTRACT INTRODUCTION Study Area Materials and methods Results and discussion REFERENCES

 

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