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Wetland Biogeochmistry Laboratory, University of Florida, 106 Newell Hall, P.O. Box 110510, Gainesville, FL 32611
* Corresponding author (krr{at}ufl.edu)
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ABSTRACT |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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A major limiting factor of microbial growth in short term studies is the utilization of readily degradable compounds of the dissolved organic carbon (DOC) pool (Hoppe, 1983). Short term studies primarily determine microbial respiration on the basis of the DOC pool. In long term studies, utilizable portions of the DOC pool are depleted, and heterotrophic microbial activity measurements are often based on utilization of large organic compounds which must be acted upon by extracellular enzymes, resulting in lower microbial activity (Chrost, 1991).
Organic matter degradation in wetlands is often limited by the availability of electron acceptors rather than electron donors (Reddy and D'Angelo, 1994; Amador and Jones, 1995; McLatchey and Reddy, 1998). Oxygen is the most important electron acceptor in terrestrial systems, but is usually limited to the upper soil surface and the overlying water column in wetlands. A sequential reduction of electron acceptors with depth in soils generally proceeds in the order of O-2 > NO-3 > SO2-4 > HCO-3 on the basis of theoretical thermodynamic energy yields to microorganisms (Billen, 1982; Reddy and D'Angelo, 1994). Thus, degradation rates of DOC in wetland soils are higher under drained conditions with O2 as the primary electron acceptor, and generally decrease in anaerobic environments depending on the availability of alternate electron acceptors (Lovley and Klug, 1986; McLatchey and Reddy, 1998; D'Angelo and Reddy, 1999).
The objectives of this study were to determine (i) rates of potential heterotrophic microbial activities (represented by CO2 and CH4 production rates) under drained (aerobic) and flooded (anaerobic) conditions, and (ii) the influence of added substrates and electron acceptors on heterotrophic microbial activities.
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MATERIALS AND METHODS |
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Detritus and Soil Sampling and Characterization
Detritus and soil samples were collected from eight stations along the P gradient south from the primary point of inflow water (S10-C) to WCA-2a (Table 1). Sampling stations encompassed low and high concentrations of soil P (approximate range of 400–2000 mg P kg-1 soil) and three vegetative zones (Typha, mixed transitional area, and Cladium/sloughs). Detritus and soils were collected during August 1996 and March 1997 to represent the seasonal wet and dry seasons. Samples consisted of recently deposited, readily distinguishable plant detritus on the soil surface and two soil depth intervals. Detritus was collected by hand at each of the sampling stations from above the cored soil. Soil cores were obtained by driving an aluminum corer (i.d. = 14.6 cm) to a depth of approximately 40 cm. At each sampling station, four soil cores were obtained 1–2 m apart. Each soil core was sectioned into 0–10 and 10–30 cm layers. Respective layers of all four cores were combined into one bulk sample and homogenized for use in experiments. A portion of samples was immediately used in field incubations as described below, while remaining samples were placed in airtight bags and stored on ice for use in laboratory experiments. Subsequently, all samples were stored at 4°C until analysis.
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Influence of Drained and Flooded Conditions on Heterotrophic Microbial Activity
These experiments were designed to provide an estimate of in situ heterotrophic microbial activity (CO2 production rates). Hence, experiments were initiated within 2 hr of sample collection and samples were incubated on site in floodwater. To provide for estimate of field CO2 production rates, a short term incubation was utilized. On the basis of preliminary studies (data not presented), CO2 production rates were linear over an 8 hr incubation period, so a 4 hr incubation period was selected.
Treatments included measurement of CO2 production under drained and flooded soil conditions. For drained conditions, detritus and soil were placed onto glass fiber filters and excess water was drained for approximately 5 min. For flooded conditions, incubations were carried out using field wet soil samples. Incubations were done in triplicate and with controls to account for background CO2 concentrations.
Wet Season (August 1996)
This field study involved the measurement of CO2 production under drained and flooded soil conditions. Schott media bottles with screw-type lids containing 10 g drained, moist soil and a NaOH trap were sealed under an atmosphere of 21% O2 to facilitate aerobic conditions. A flooded (anaerobic) treatment containing 10 g wet samples of flooded soil with a N2 headspace was also included. For substrate induced respiration (SIR) measurements, both drained and flooded treatments were supplemented with glucose at an excess concentration of 25 mg C g soil -1 based on results of previous experiments (DeBusk, 1996). Incubations were carried out at ambient floodwater temperature (28 ± 2°C).
Carbon dioxide production was quantified using an alkali trap method in which 10 mL of 0.1–0.2 M NaOH was added to small vials placed into incubation bottles. After 4 hr of incubation, vials containing NaOH traps were removed and capped. The CO2 trapped in NaOH was subsequently analyzed by titration with acid (Zibilske, 1994). For titrations, BaCl2 was added to NaOH traps and remaining NaOH was titrated with 0.05–0.1 M HCl to the phenolphthalein endpoint. At the end of incubation, detritus and soil samples were oven-dried at 70°C to determine sample dry weight per incubation bottle.
Dry Season (March 1997)
The experimental design involved measurements taken within 2 hr of sample collection (basal CO2 production) and a later incubation with added substrates (SIR) initiated approximately 6 hr after sample collection. Incubations for basal CO2 production were carried out at ambient floodwater temperature (28°C) for a period of 4 hr. The experimental design was similar to that described for the wet season experiment.
Following measurement of basal CO2 production, additional incubations were initiated on the same samples to determine SIR. Substrate sources included either a mixture of C sources (glucose, citrate, malate, oxalate, and acetate; each component contributing 20% of the total C in the mixture) or a C, N, S, and P source (peptone) at rates of 25 mg C g soil-1. Both substrate solutions were buffered to pH 7 and autoclaved. After completion of field incubations, substrate solutions was added to incubation bottles, a NaOH trap (3 mL of 0.2 M) was then added, and samples were either incubated under an atmosphere of 21% O2 or were purged with N2. After a 4 hr incubation at 28°C, NaOH traps were removed and sealed with caps fitted with rubber septa.
Unlike the previous study during the wet season, CO2 trapped in NaOH was determined by gas chromatography after acidification of NaOH. This method was found to be more sensitive to levels of CO2 than the titration method. To the enclosed NaOH in vials, excess HCl was added through septa to neutralize NaOH and result in a pH of below 2.0. A portion of the resulting CO2 in the headspace was sampled by syringe for injection to the gas chromatograph. Carbon dioxide was measured using a Shimadzu GC-8A gas chromatograph fitted with a thermal conductivity detector (30°C), He as carrier gas, and a 0.3 cm by 2 m Poropak N column (Supelco Inc., Bellefonte, PA) at 25°C. Gas pressure in the vials was measured using a digital pressure meter (Kane-May, UK). Carbon dioxide trapped in NaOH was then calculated by a modification of the method described by Martens (1987) in which the universal gas law, pressure in the vials, solution pH, CO2 concentration in the headspace, and solution and headspace volumes were used to calculate trapped CO2.
Influence of Added Inorganic Electron Acceptors on Heterotrophic Microbial Activity
This experiment utilized detritus, 0–10 cm, and 10–30 cm soil sampled from 8 stations along the P gradient in WCA-2a in March 1997 (dry season). Four treatments (6 replicates per treatment) were applied to each soil depth at each station. Treatments included incubation under aerobic, nitrate reducing, sulfate reducing, and methanogenic conditions. The electron acceptors, NO-3 and SO2-4, were added at an electron equivalent basis and at concentrations determined in preliminary experiments to be in excess of the concentrations needed. A KNO3 solution was added to soil resulting in an initial concentration of 1600 µg NO3–N g-1 dry soil. A K2SO4 solution was added to soil resulting in initial concentration of 2300 µg SO4-S g-1 dry soil.
Approximately 10 g wet soil was added to incubation bottles followed by addition of electron acceptors. For O2 treatments, samples were drained on glass fiber filters to remove excess water and to allow for aerobic conditions. A vial containing 5 mL of 0.1 M NaOH was placed inside incubation bottles to trap evolved CO2. Bottles were purged with N2 gas (99.9% purity) for all treatments with the exception of the O2 treatment. For the O-2, NO-3, and SO2-4 reducing treatments, the NaOH traps were removed from incubation bottles at 4, 8, 16, and 24 hr and then at approximately 2 d intervals up to 10 d. For the methanogenic CO2 production treatments, NaOH traps were sampled at 4, 8, 16, and 24 hr and then periodically up to 40 d. At the end of each sampling period, bottles from anaerobic treatments were purged with N2. The NaOH traps were analyzed for CO2 by gas chromatography after acidification. Methane accumulation in the headspace was monitored by periodic sample headspace injection into a Shimadzu gas chromatograph-8A fitted with flame ionization detector (160°C), N2 as the carrier gas, and a 0.3 cm by 2 m Carboxyn 1000 column (Supelco Inc., Bellefonte, PA) at 160°C. Controls were included to account for background CO2 and CH4.
Data Analyses
Comparison of treatments and significant differences were determined using one-way and three-way Analysis of Variance (ANOVAs) with a Fisher's LSD at P < 0.05 (CoStat, Minneapolis, MN). A completely randomized experimental design was utilized with factors being electron acceptor additions, stations, soil depth, or season. Correlation coefficients were determined using CoSTAT (Minneapolis, MN). Field replications were developed by combining adjacent sampling sites into distinct groups on the basis of soil P concentrations and vegetation type. The groupings for site comparisons were: stations 1–3 (P impacted area growing Typha), stations 4–6 (transitional area having both Typha and Cladium), and stations 7–8 (P unimpacted area growing Cladium). Statistical differences at P < 0.05 were compared between the P impacted, transitional, and unimpacted areas.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Influence of Drained and Flooded Conditions on Heterotrophic Microbial Activity: Dry Season
Basal CO2 production rates were highest in detritus and significantly (P < 0.05) decreased with depth under both drained and flooded conditions (Fig. 1). Production rates in drained samples were greater (P < 0.05) than rates in flooded samples for both detritus and 0–10 cm soil. However, no difference in CO2 production rates between drained and flooded conditions was observed in the 10–30 cm depth. Both drained and flooded basal CO2 production rates significantly (P < 0.05) decreased along the P gradient in the detritus and 0–10 cm depth but not in the 10–30 cm depth.
Under drained conditions, addition of peptone increased CO2 production rates more than the mixture of C sources in the detritus (P < 0.05) (Table 3). Production rates of CO2 under drained conditions was highest in the detritus and decreased with depth regardless of which substrates were added. Under flooded conditions, addition of peptone increased CO2 production more than addition of C sources but only in the detritus (P < 0.05). Flooded soil CO2 production rates were highest in the detritus and decreased with depth regardless of which substrate was added. Drained soil CO2 production rates were greater than flooded rates in detritus and 0–10 cm layers with either substrate added. No seasonal differences in CO2 production rates were observed in detritus and soil.
Basal CO2 production rates measured during the wet and dry season experiments were significantly (P < 0.05) related to the total P content of detritus and soil (Fig. 2) . Anaerobic CO2 production rates were approximately 64% of the aerobic rates (Fig. 3) .
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Heterotrophic microbial activity was generally highest at the P impacted area and decreased at unimpacted areas (Fig. 4 and 5) . Differences among various electron acceptor treatments also were evident. Aerobic respiration was significantly (P < 0.05) higher than all other modes of respiration and methanogenesis. Differences with soil depth were also observed, with CO2 and CH4 production rates being higher in detritus than in underlying soil.
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Soil depth had a strong effect on both k1 and k2 values. Under aerobic, nitrate reducing, sulfate reducing, and methanogenic CO2 producing conditions, both k1 and k2 were significantly (P < 0.05) highest in detritus and decreased with depth (Table 4). However, CH4 production exhibited slightly different relationships with depth. Although CH4 production rates in k1 and k2 were significantly highest in detritus, there were no differences in k1 and k2 values between 0–10 cm and 10–30 cm soil.
The k1 and k2 values under aerobic conditions was significantly (P < 0.05) greater than k1 and k2 for all other electron acceptor treatments in detritus, 0–10 cm, and 10–30 cm soil (Table 4). The k1 values for CH4 production were significantly (P < 0.05) lower than values for all other treatments in detritus and soil. The k1 values under denitrifying and sulfate reducing conditions were similar in detritus but k1 values were significantly (P < 0.05) higher under sulfate reducing conditions than under denitrifying conditions in 0–10 cm soil.
For k2 values, no differences between sulfate and nitrate reducing conditions were observed in any soil depth. However, k2 values under nitrate and sulfate reducing conditions were significantly greater than methanogenic CO2 and CH4 producing conditions at all soil depths. The k1 and k2 values for aerobic conditions were plotted against values for other electron acceptor treatments (Fig. 6 and 7) , with CO2 production under nitrate reducing conditions being 30% of aerobic production for k1, but increasing to 42% for the k2 phase. The sulfate reduction rate was 44% of the aerobic rate in k1 phase but decreased to 29% in the k2 phase. The CH4 production rate was only up to 9% of the aerobic CO2 production rate.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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Basal and substrate induced CO2 production rates decreased with depth in the soil profile, also corresponding to decreases in extracellular enzyme activity and substrate quality with depth (DeBusk and Reddy, 1998; Wright and Reddy, 2001). Many factors, such as dissolved O2, enzyme activity, electron acceptor concentrations, and substrate quality also decreased with soil depth (McKinley and Vestal, 1992; D'Angelo and Reddy, 1994; DeBusk, 1996). Substrate quality can be measured on the basis of on a lignocellulose index (LCI), which is a ratio of lignin to lignocellulose (Melillo et al., 1989). Lignin content in Everglades soils increased with depth in conjunction with decreases in cellulose content: thus substrate quality decreases with depth and has been implicated in regulating organic matter degradation (Swift et al., 1979; Benner et al., 1984; DeBusk, and Reddy, 1998). Soil depth had a strong effect on both k1 and k2 values in our laboratory studies. Under aerobic, nitrate reducing, sulfate reducing, and methanogenic CO2 producing conditions, both k1 and k2 were significantly (P < 0.05) highest in detritus and decreased with depth (Table 4). Thus, substrate quality may be a primary controlling factor of heterotrophic microbial activities in subsurface soils.
Oxygen appeared to regulate organic matter degradation in these studies, with CO2 production rates for drained soil being higher than for flooded soil. This was expected as microbial activity is normally greater under aerobic conditions (Benner et al., 1984; Bridgham and Richardson, 1992; DeBusk and Reddy, 1998; McLatchey and Reddy, 1998; D'Angelo and Reddy, 1999). In wetland soils, degradation generally proceeds on the basis of a thermodynamically favored sequential reduction of electron acceptors O-2, NO-3, SO2-4, and HCO-3 (Zehnder and Stumm, 1988; Reddy and D'Angelo, 1994). Based on the theory of preferential use of electron acceptors by microorganisms, aerobic microbial activities should be greater than anaerobic activities since more energy can be obtained by microorganisms using O2. In the short term incubation field experiments, the anaerobic CO2 production rate was 64% of the aerobic rate (Fig. 3), which was somewhat higher than other rates measured under laboratory conditions (Bridgham and Richardson, 1992; DeBusk and Reddy, 1998). In our short term incubation of 4 hr, O2 contamination from soil mixing may have contributed to an inflated anaerobic CO2 production rate. DeBusk and Reddy (1998) reported that anaerobic CO2 production rates were 32% of the aerobic rates along the same P gradient in WCA-2a. Similar results have been reported to range from 34–63% (Bridgham and Richardson, 1992) and 37% (Benner et al., 1984). Soils along the P gradient often have low NO-3 concentrations, thus, upon initiation of experiment, population sizes of denitrifiers were likely low. After an adjustment period during the k1 phase, nitrate reducers likely increased in population size, thus increasing CO2 production up to 42% of aerobic CO2 production rates during the k2 phase.
In our experiments, the sequential reduction of electron acceptors held true for O2 (highest rates) and HCO-3 (lowest rates) but not for NO-3 and SO2-4. In most cases, there were no differences in heterotrophic microbial activities under nitrate and sulfate reducing conditions, perhaps due to the presence of SO2-4 and a general lack of NO-3 in soils along most of the P gradient. It was expected that CO2 production rates would be greater under nitrate rather than sulfate reducing conditions. Perhaps relatively higher SO2-4 concentrations and lower NO-3 concentrations in soils along the P gradient tended to support sulfate reduction and limit denitrification. The decrease in sulfate reduction rates along the P gradient in both the rapid k1 phase and more stable k2 phase may be due to P loading or the presence of high SO2-4 concentrations near the inflow point. Sulfate concentrations have been detected at high levels along the P gradient (Schipper and Reddy, 1995), and sulfate reducing bacteria have been shown to be enhanced by P (Drake et al., 1996).
Addition of C substrates in field experiments tended to increase CO2 production rates but only for the detritus. Addition of peptone increased CO2 production in several cases compared with addition of a C source only. Peptone is a mixture of various compounds containing C, N, P, S, and micronutrients, so it appears that addition of nutrients other than C stimulated CO2 production. Since peptone contains both N and P, either or both of these nutrients may be responsible for increased CO2 production compared with treatments receiving C sources only. However, since P concentrations were already high at impacted areas, and since peptone increased CO2 production rates in these areas, it seems unlikely that additional P in the form of peptone would be responsible for increased CO2 production rates. Amador and Jones (1995) showed that additions of C and P increased C mineralization rates in P limited areas of the Everglades but not in areas of P enrichment. The N supplied by peptone may be responsible for the increased respiration rates observed in P impacted areas. Indeed, N may become limiting to heterotrophic microbial activity in areas of high P concentrations (DeBusk, 1996).
Heterotrophic microbial activity measured in field and laboratory experiments utilizing various electron acceptors was significantly (P < 0.05) correlated with many soil physical and chemical parameters, including soil total P, labile C, extractable C, and microbial biomass C. Heterotrophic microbial activity has been reported in several studies to be enhanced by P additions or in high P soils (Amador and Jones, 1993; Bridgham and Richardson, 1992; Amador and Jones, 1995; DeBusk and Reddy, 1998). Labile and extractable C represent the most utilizable portions of soil total C, so they were expected to be related to CO2 and CH4 production rates. However, soil total C was not related to heterotrophic microbial activity, suggesting that the bulk of soil total C was not utilizable. Carbon dioxide production rates in drained detritus and soil were significantly (P < 0.05) correlated with b-d-glucosidase activity (Wright and Reddy, 2001). The significant relationships between enzyme activity and heterotrophic microbial activity in drained soil suggests that enzyme activity is closely coupled with microbial CO2 and CH4 production in Everglades soil, most likely in the longer term k2 phase. However, anaerobic CO2 production rates and enzyme activities were not significantly correlated.
Heterotrophic microbial activity was significantly enhanced by the draining of soil and exposure to O2. This has important implications with regard to water management, organic matter degradation, and nutrient cycling. Exposure of soil to O2, as it occurs in periods of low rainfall or low water inputs, increases heterotrophic microbial activity. This increase in microbial activity contributes to enhanced organic matter degradation rates and increased regeneration and cycling of nutrients in wetlands which could potentially lead to increased nutrient concentrations in the water column.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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ACKNOWLEDGMENTS |
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Received for publication August 22, 2000.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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