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Wetland Biogeochemistry Lab., Soil and Water Science Dep., 106 Newell Hall, Univ. of Florida, Gainesville, FL 32611
Corresponding author (jrwh{at}gnv.ifas.ufl.edu)
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Abbreviations: ANOVA, analysis of variance • MBC, microbial biomass C • MBN, microbial biomass N • SOD, soil oxygen demand • WCA, Water Conservation Area
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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The redox status of a wetland soil system can exert substantial control over the cycling of N (Reddy and Patrick, 1975). Mineralization of organic N can proceed under both aerobic and anaerobic conditions. Due to the restricted supply of O2 in wetland soils, the influence of alternate electron acceptors on microbial catabolic processes can mediate the rate at which organic matter decomposition occurs in wetland soils (McLatchey and Reddy, 1998; D'Angelo and Reddy, 1999). The concept of free energy determines the order by which electron acceptors are used by microbial populations, in the order: O2, NO-3, SO2-4, and finally CO2 reduction. The composition of the functional microbial pool and rates of organic N mineralization will, therefore, vary under dominance of any single inorganic electron acceptor.
Specific functional microbial communities become established in the soil profile, dependent on aeration status and the availability of inorganic electron acceptors. Aerobes are found within the surface layer. The NO-3 reducers are located below these in the profile, with SO2-4 reducers and methanogens located deepest in the soil profile and out of range of influence of O2 (Reddy and D'Angelo, 1994). This layer cake model of electron acceptor consumption has also been demonstrated for a coastal sediment (Sorensen et al., 1979). An allochthonous supply of inorganic electron acceptors from either surface or groundwater sources could significantly alter soil nutrient dynamics and wetland function.
Differences in net N mineralization can be investigated during short time periods (h) with simple substrates (amino acids) because the rate limiting steps of soil organic matter breakdown and decomposition have been removed by providing a readily hydrolyzable substrate. The C/N ratio of various amino acids is much lower than the C/N ratio of the microbial pool resulting in substantial net N release (Alef and Kleiner, 1986). Amino acid utilization as a respiratory substrate (electron donor) provides a measure of the activity of the heterotrophic microbial population (Alef et al., 1988; Hopkins et al., 1993, 1997). Substrate-induced N mineralization is linear in the short term (a few hours) and has been shown to occur with minimal or no lag phase in soil (Alef and Kleiner, 1986). This method measures the presently active soil microbial population responsible for the final step of organic N mineralization, deamination, without allowing sufficient time for significant turnover and de novo biomass synthesis (Hopkins and Ferguson, 1994; Franzluebbers et al., 1996; McLatchey and Reddy, 1998).
The Florida Everglades have been affected by nutrient loading from urban and agricultural surface water runoff, in particular, in the Water Conservation Areas (WCA) (DeBusk et al., 1994). Specifically, WCA-2A has been receiving nutrient-laden (N and P) drainage waters for the past 40 yr (Koch-Rose et al., 1994). Peat accretion rates have increased in areas receiving surface agricultural and urban drainage water (Koch and Reddy, 1992; Craft and Richardson, 1993; Reddy et al., 1993). The nutrient loading has been documented in the spatial distribution of surface soil total P in this historically oligotrophic wetland system. Surface soil total P concentrations range from highs of 
1600 mg kg-1 at the surface water inflow points to a background concentration of 450 mg kg-1 in unimpacted areas (Reddy et al., 1993; DeBusk et al., 1994; Reddy et al., 1999). A gradient of N and P in the water column and periphyton (algal) tissue has also been documented along the same eutrophic transect in WCA-2A (McCormick and O'Dell, 1996). The vegetation community began a shift from a dominant sawgrass (Cladium jamaicense Crantz) marsh (Davis, 1943) towards a dominant cattail (Typha domingensis Pers.) vegetative community proximal to all surface water inflow points (Davis, 1991; Craft and Richardson, 1997).
The hypothesis tested was that N mineralization rates of organic matter would be affected by the dominance of selected inorganic electron acceptors. The objectives of this study were to determine (i) the rates of potential mineralization of native organic N associated with detritus and soil under conditions of various inorganic electron acceptors dominance (aerobic, NO-3, and SO2-4 reducing, and methanogenic conditions) and (ii) the relationship between easily measured soil characteristics (including total P) and potential organic N mineralization rates.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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10 km into the natural (unimpacted, with respect to P concentrations) marsh characterized by stunted stands of sawgrass (Cladium spp.), separated by shallow sloughs and dominated by floating and attached cyanobacterial mats. Sampling stations were located at distances of 1.4, 2.3, 3.3, 4.2, 5.1, 7.0, 8.4, and 10.1 km (Fig. 1). Water depths varied seasonally from <2 cm to 2 m along the transect length. Sampling along the transect was not designed to identify differences between separate stations, but rather to investigate the gradient or trends between soil characteristics and selected microbial N processes, including aerobic and anaerobic organic N mineralization.
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Samples were transferred into 2-L polyethylene containers within 24 h of collection and stored refrigerated at 4°C until subsequent characterization (within 1 mo). Samples of detritus and soil for the determination of anaerobic N mineralization rates were collected in March 1997 and samples for determination of aerobic N mineralization rates were collected in October 1997. Soils were collected at different times in order to keep refrigerated storage time to less than 1 mo.
Soil Characterization
Moisture (percentage oven dry weight basis) was determined by drying 20 g of a field-moist subsample in a forced-air drying oven at 70°C to constant weight. 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). Total P was determined using an automated ascorbic acid method (USEPA, 1983).
Soil oxygen demand (SOD) was determined for a single core collected in February 1996 at Station 1 (1.4 km from inflow). The top 36 cm of soil were sectioned into 2-cm intervals, and 10 g (wet weight) of sample were added to 300 mL of distilled, deionized water in a capped, continuously stirred BOD bottle. Dissolved O2 was monitored using a YSI Model 58 oxygen meter with probe (YSI, Yellow Springs, CO). Soil oxygen demand was calculated as the difference in O2-saturated soil/water slurry at t = 0 minus measured dissolved O2 concentrations after 8 h, divided by the dry weight of the soil sample and time elapsed between measurements (American Public Health Association, 1992).
Extractable NH+4 was determined by shaking triplicate soil samples with 25 mL of 2 M KCl at a ratio of approximately 1:40 (grams dry soil/extractant) for 1 h on a reciprocating shaker. Samples were centrifuged for 10 min and vacuum-filtered through Whatman no. 42 filter paper. The supernatant was collected and refrigerated at 4°C and NH4–N concentrations were determined colorimetrically (USEPA, 1983).
Microbial Biomass
Microbial biomass C was determined by fumigation-extraction (Vance et al., 1987). Three 5-g subsamples for each soil interval and sampling station were fumigated for 24 h, and three were not fumigated. All samples were extracted with 20 mL of 0.5 M K2SO4 and filtered through no. 42 Whatman filter paper (White and Reddy, 2000). Dissolved organic C was determined on a Dohrman DC 190 C analyzer (Dohrman, Santa Clara, CA). Microbial biomass C was determined by subtracting the extractable total organic C in the triplicate controls (nonfumigated) from the triplicate chloroform-treated samples. A combined extraction efficiency (kEC) factor of 0.37 was applied, using a previous calibration for organic soils (Sparling et al., 1990).
Microbial biomass N was determined by fumigation-extraction (Brookes et al., 1985). Ten milliliters of extract from the microbial C procedure was subjected to Kjeldahl-N digestion using the salicylic acid modification (Bremner and Mulvaney, 1982). Samples were brought to a total volume of 20 mL after digestion and transferred into 30-mL scintillation vials. Extracts were analyzed for NH4–N colorimetrically (USEPA, 1983). Microbial biomass N was determined by subtracting the extractable NH4–N of the triplicate nonfumigated samples from triplicate fumigated samples. An extraction efficiency (kEN) value of 0.54 was applied (Brookes et al., 1985).
Potential Organic Nitrogen Mineralization under Aerobic Conditions
Aerobic N mineralization rates of detritus and soil were determined in constantly stirred reactors in order to prevent denitrification losses (McLatchey and Reddy, 1998). Approximately 300 g wet weight of soils and litter were placed in triplicate 1-L Erlenmeyer glass flasks and mixed with 400 mL of water. The moisture content of the soil slurries in the reactors averaged 97%.
The contents of the flasks were continually mixed, in concert with continuous aeration with room air (using an aquarium pump connected to glass tubing inserted through a butyl rubber stopper in each flask) to maintain aerobic conditions in the slurry. The redox status of soil slurries was monitored using a Fisher brand Accumet 1002 combination electrode with platinum band (Fisher Scientific, Pittsburgh, PA), and temperatures were recorded with mercury thermometers. The average temperature of the reactors was 30°C. Reactors were covered with opaque paper to shield the soil from direct light.
Ten milliliters of slurry was collected from each reactor daily for 15 d, extracted with 10 mL of 2 M KCl, and filtered through Whatman no. 42 filter paper. The supernatant was refrigerated at 4°C for subsequent, automated, colorimetric analysis of NH+4 and NO-3 (USEPA, 1983).
Aerobic N mineralization rates were determined by summing the inorganic N at each sampling time (NH+4 + NO-3) and fitting a regression line through the data. Rates were expressed in units of milligrams N per kilogram dry weight of soil per day.
Organic Nitrogen Mineralization under Anaerobic Conditions
The organic N mineralization rate of soils and detritus under dominant NO-3 reducing, SO2-4 reducing, and methanogenic conditions were determined for the March 1997 samples. Triplicate glass serum bottles were prepared by adding 5 g of moist soil and 5 mL of distilled, deionized water. Moisture content of the soil slurries averaged 97%. Bottles were capped and sealed with aluminum crimps. The headspace was evacuated to -85 kPa and replaced with 99.99% O2-free N2 gas. For the respective treatments, NO-3 as KNO3 and SO2-4 as K2SO4 were added on an electron equivalent basis and at levels determined in preliminary investigations to be in excess, assuming a 15-d incubation period for a 10-g wet weight sample. One-half milliliter each of the respective solutions was applied to the individual treatments at the start of the incubation and again at t = 8 d. The concentrations of electron acceptors in the sample soil solution at the time of spiking were 80 mg NO3–N L-1 and 115 mg SO4–S L-1 for the NO-3 and SO2-4 reducing conditions, respectively. Serum bottles were incubated in the dark at 30°C for 15 d and were shaken by hand for 30 s d-1 during the length of the incubation to overcome diffusion constraints. A set of triplicate time-zero controls was extracted with 2 M KCl at the start of the incubation period.
In preparation for incubation under methanogenic conditions, detritus and soil samples (including the controls) were preincubated in the dark under anaerobic conditions at 30°C. Headspace gas was periodically monitored by collection of a 50-µL gas sample. Methane concentrations were determined using a Shimadzu 8 AIF gas chromatograph (Shimadzu, Kyoto, Japan) equipped with a flame ionization detector (110°C) with N2 as the carrier gas and a stainless steel Carboxen 1000 column (Supelco, Bellefonte, PA) maintained at 160°C in order to determine the onset of CH4 production. The preincubation period (length of time the controls were incubated until CH4 appeared) averaged 4 d for detritus and the 0- to 10-cm soil depth, while averaging 9 d for the 10- to 30-cm soil interval. Preliminary investigations indicated that the preincubation period was not required for the samples under NO-3 reducing and SO2-4 reducing conditions as both electron acceptors were being consumed within 1 h from the time of spiking, demonstrating active microbial use.
All samples were extracted with 30 mL of 2 M KCl at the terminus of the incubation period (15 d). Samples were filtered through Whatman no. 42 filter paper, collected in 25-mL scintillation vials and refrigerated at 4°C for subsequent automated, colorimetric analysis of NH+4 (USEPA, 1983).
Substrate Induced Nitrogen Mineralization
Soil and detritus samples were prepared in the same manner as described for the anaerobic mineralization study. Sample bottles were spiked with NO-3, SO2-4, or no additions and were incubated for 15 d in order to assess microbial activity under NO-3 reducing, SO2-4 reducing, and methanogenic conditions, respectively. To triplicate subsamples, 0.5 mL of solution containing 200 mg L-alanine (C3H7NO2)-N L-1 was added and incubated in the dark at 30°C. Samples were extracted at the end of the 4-h incubation with 30 mL of 2 M KCl and filtered through no. 42 Whatman filter paper. The supernatant was stored refrigerated at 4°C until subsequent automated colorimetric analysis for NH4–N (USEPA, 1983). The substrate-induced N mineralization rate (mg N kg-1 h-1) was calculated as the difference in extractable NH+4 between the spiked incubation and controls samples divided by the 4-h incubation period and dry weight of the sample.
Data Analysis
Soil characteristics and microbial processes were statistically related using Pearson's product-moment correlation and regression analysis. All data were checked for homogeneity of variances and log transformed prior to statistical comparisons when appropriate. Analysis of variance (ANOVA) and Fisher's least significant difference tests were used to make comparisons between treatments after pooling data from all stations at each sampling depth using the StatGraphics software program (Manugistics, Rockville, MD).
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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The results of the SOD measurements on soil samples demonstrated high O2 consumption rates in the surface soil that decreased exponentially with depth (Fig. 2) . The decrease in SOD with depth was attributed to lower microbial activity probably because of fewer readily available C compounds at lower depths (DeBusk, 1996). Oxygen, used by aerobic heterotrophs as the terminal electron acceptor, is consumed rapidly during microbial respiration in the surface soil. The high oxygen demand of the surface soil prevents O2 from diffusing downward from the water column into the sediments, assuring the bulk of the wetland soil remains anaerobic. In addition, the high O2 consumption rate in the soil profile underscores the importance that alternate inorganic electron acceptors may have in mediating the mineralization of organic N in wetlands.
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Microbial Biomass
Microbial biomass C and N were correlated (P < 0.01) with total P (r = 0.73 and 0.67, respectively; Table 2). Both microbial biomass C and N were also significantly negatively correlated with depth (r = -0.081 and -0.74, respectively; Table 2). Microbial biomass C and N were positively correlated (r = 0.88; P < 0.01) with one other. The best-fit linear model of microbial C and N yielded an average C/N ratio of 12.9 for the microbial pool.
Microbial biomass C and N were both positively correlated (P < 0.01) with extractable NH+4 (r = 0.89 and 0.95, respectively; Table 2). Extractable NH+4 appears to be a useful indicator of the size of microbial pool size measured by chloroform-fumigation extraction with NH+4 concentrations describing 79 and 90% of the variability of microbial biomass C and N, respectively.
Organic Nitrogen Mineralization Rates
Significant differences in organic N mineralization rates of native organic matter with depth were seen under each of the aerobic, NO-3 reducing, SO2-4 reducing, and methanogenic conditions. Under aerobic conditions, organic N mineralization rates averaged 237, 143, and 75 mg N kg-1 d-1 for detritus, 0- to 10-, and 10- to 30-cm soil depths, respectively (Table 3). Rates of N mineralization with depth also followed a similar pattern for NO-3 reducing, SO2-4 reducing, and methanogenic conditions with significantly higher rates in the detritus, significantly lower rates in the 0- to 10-cm soil depth and lowest rates found in the 10- to 30-cm soil depth. These results suggest that the surficial detritus provides the greatest source of inorganic N among the soil compartments sampled. Previous studies of flooded peat soils have also documented decreased organic N mineralization with increasing soil depths (Franzluebbers et al., 1995; Hossain et al., 1995).
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Similar results have been observed for another organic rich Florida wetland soil where all soils were placed in constantly stirred reactors (McLatchey and Reddy, 1998). A survey of 10 wetland soils from throughout the USA found that C mineralization rates were three times greater than rates under anaerobic conditions for surface soils in constantly shaken vials (D'Angelo and Reddy, 1999). The results of these studies suggest that the difference in methodology (constantly stirred vs. daily short-term mixing) did not dramatically alter our results, as we found similar N mineralization rates difference for surface wetland soil samples. In addition, these Everglades organic soils contain high moisture contents (90–95%) and low mineral matter. Therefore, the continual stirring would not substantially break up the soil further, as the soil was already in a slurry at 97% moisture contents.
Aerobic N mineralization rates were significantly correlated with rates under NO-3 reducing (r = 0.59), SO2-4 reducing (r = 0.55), and methanogenic (r = 0.81) conditions at P < 0.01, respectively (Table 4). Standard correlation procedures use mathematically equivalent calculations for calculating coefficients of determination based on linear relationships. The coefficient of determination (R2) improved to 0.54 and 0.47 for mineralization under NO-3 and SO2-4 reducing conditions, respectively, compared with aerobic conditions when employing a nonlinear equation (Fig. 3) . Comparisons of aerobic N mineralization rates of soil and detritus with rates under methanogenic conditions were best described by a linear regression. These results point out the existence of a variety of functional microbial groups viable in the soil (Drake et al., 1996), which can play a significant role in organic N mineralization in wetland soils.
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Rates under NO-3 reducing conditions were significantly correlated with total P (r = 0.53), MBC (r = 0.70), MBN (r = 0.80), and extractable NH+4 (r = 0.85) at P < 0.01. Extractable NH+4 was again the strongest indicator of organic N mineralization rates, with the equation of the best-fit line explaining 72% of the variability. A similar relationship was seen under SO2-4 reducing conditions, with N mineralization rates correlated with total P, MBC, MBN, and extractable NH+4 (Table 4). Extractable NH+4 was the strongest indicator of N mineralization rates.
The N mineralization rates under methanogenic conditions were the most strongly correlated rates with measured soil characteristics, including total P (r = 0.60), MBC (r = 0.83), MBN (r = 0.90), and extractable NH+4 (r = 0.92). Again, extractable NH+4 was the strongest indicator of potential N mineralization rates.
These results suggest that for the waterlogged, highly reduced organic soils of WCA-2A, extractable NH+4 was an excellent indicator of potential organic N mineralization of soil and detritus under aerobic, NO-3 reducing, SO2-4 reducing, and methanogenic conditions. In addition, mineralization rates were strongly correlated with the size of the microbial pool, suggesting microbial pool size was a relatively sensitive indicator of the heterotrophic microbial pool activity and, consequently, N cycling.
Substrate-Induced Nitrogen Mineralization
Substrate-induced N mineralization was measured under anaerobic conditions to determine the relative activity of the portion of the microbial pool responsible for the final steps in organic N mineralization (deamination) under NO-3 reducing, SO2-4 reducing, and methanogenic conditions. Substrate-induced N mineralization rates under NO-3 reducing and methanogenic conditions were significantly correlated with distance from the inflow. Both substrate-induced N mineralization rates under NO-3 and SO2-4 reducing conditions, r = -0.42 and -0.44, respectively, were negatively correlated (P < 0.05) with depth, and substrate-induced N mineralization rates under methanogenic conditions were weakly negatively correlated with depth (P < 0.10).
There were no differences in substrate-induced N mineralization among anaerobic electron acceptor conditions (Table 5). In order to remove N in the structure of L-alanine, a simple deamination is required. It is likely that the effect of electron acceptors on organic N mineralization is linked only to the return of energy in breaking the C-N bonds in the larger organic molecules, as was the case in mineralization of the larger, more complex native organic matter molecules. Deamination was not the rate-limiting step in the breakdown of organic N as substrate-induced N mineralization rates were much larger (Table 5) than N mineralization rates of the native organic matter (Table 3). Rates were not affected by dominance of any particular electron acceptor. Substrate-induced N mineralization rates, under NO-3 reducing conditions only, were significantly correlated with MBN. It has been shown that the presence and activity of extracellular enzymes can be more significant than measured soil characteristics in regulating N release from amino acids in these soils (McLatchey and Reddy, 1998). Therefore, substrate-induced N mineralization did not provide any useful indication of the relative rates of organic N mineralization under aerobic, NO-3 reducing, SO2-4 reducing, and methanogenic conditions.
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2 km from the inflow points, and then only to the detrital and 0- to 10-cm soil depths (White and Reddy, 1999). Consequently, organic N mineralization under NO-3 reducing conditions is restricted to areas along the transect proximal to the surface water inflow points. The concentrations of SO2-4 are also highest at the inflow points (Koch-Rose et al., 1994). Therefore, the cattail areas are likely to undergo relatively high organic N mineralization rates in situ due to the availability of O2 part of the year and a nearly continuous supply of NO-3 and SO2-4 during the summer months when hydraulic loading is highest.
Soils in the natural sawgrass marsh, located in the interior of WCA-2A, exhibit very low denitrifying enzyme activity, which suggests very little allochthonous NO-3 is present (White and Reddy, 1999). The porewater SO2-4 concentrations are also significantly lower in the sawgrass areas than in the cattail areas (Koch-Rose et al., 1994). The presence of open water dominated by periphyton, however, provides a mechanism of transporting O2 into the soil to stimulate inorganic N release within these areas. During the daylight hours, photosynthetic activity is high and redox profiles have detected aerobic conditions down into the detrital layer and even reaching the surface of the peat soil (DeBusk, 1996). This cyclic mechanism occurs daily, with low bottom water O2 levels experienced during the night. Oxidation of the detrital material might provide an important inorganic N source for macrophytes and the microbial pool, and is likely to be partly responsible for the low (0.1 cm yr-1) organic soil accretion rates in the low P sawgrass areas (Reddy et al., 1993).
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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The dominance of inorganic electron acceptors was found to have a significant influence on organic N mineralization rates of native soil organic matter. Rates of N mineralization in soil ranged from high rates under aerobic conditions to significantly lower rates under NO-3 and SO2-4 reducing conditions, and lowest under methanogenic conditions for soil and detritus. There appeared to be no effect of electron acceptor dominance on substrate-induced N mineralization rates in soil and detritus.
These data on potential rates of organic N mineralization can be used to determine the potential effect of increased loading of dissolved NO-3 and SO2-4 on the biogeochemical cycling of N in these soils. A decrease in surface water hydraulic loading to WCA-2A could potentially increase organic N mineralization rates by exposing the soil surface to O2. In addition, increased mass loading rates of NO-3 and SO2-4 in the surface could potentially increase organic N mineralization rates, thereby altering the overall balance of N in this 44000-ha wetland.
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ACKNOWLEDGMENTS |
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Received for publication August 23, 1999.
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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