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TECHNICAL REPORTS

Soil Electrical Conductivity and Water Content Affect Nitrous Oxide and Carbon Dioxide Emissions in Intensively Managed Soils

^a Department of Agronomy and Horticulture, University of Nebraska, P.O. Box 830915, Lincoln, NE 68586-0915
^b USDA-ARS Soil & Water Conservation Research Unit, Department of Agronomy and Horticulture, 116 Keim Hall, University of Nebraska, Lincoln, NE 68583-0934

^* Corresponding author (adobermann2{at}unl.edu)

Received for publication March 15, 2006.

	ABSTRACT

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Accumulation of soluble salts resulting from fertilizer N may affect microbial production of N₂O and CO₂ in soils. This study was conducted to determine the effects of electrical conductivity (EC) and water content on N₂O and CO₂ production in five soils under intensive cropping. Surface soils from maize fields were washed, repacked and brought to 60% or 90% water-filled pore space (WFPS). Salt mixtures were added to achieve an initial in situ soil EC of 0.5, 1.0, 1.5 and 2.0 dS m^–1. The soil cores were incubated at 25°C for 10 d. Average CO₂ production decreased with increasing EC at both soil water contents, indicating a general reduction in microbial respiration with increasing EC. Average cumulative N₂O production at 60% WFPS decreased from 2.0 mg N₂O-N m^–2 at an initial EC of 0.5 dS m^–1 to 0.86 mg N₂O-N m^–2 at 2.0 dS m^–1. At 90% WFPS, N₂O production was 2 to 40 times greater than that at 60% WFPS and maximum N₂O losses occurred at the highest EC level of 2.0 dS m^–1. Differences in the magnitude of gas emissions at varying WFPS were due to available substrate N and the predominance of nitrification under aerobic conditions (60% WFPS) and denitrification when oxygen was limited (90% WFPS). Differences in gas emissions at varying soil EC may be due to changes in mechanisms of adjustment to salt stress and ion toxicities by microbial communities. Direct effects of EC on microbial respiration and N₂O emissions need to be accounted for in ecosystems models for predicting soil greenhouse gas emissions.

Abbreviations: BD, Bulk density • EC, Soil electrical conductivity • PAS, Infrared photoacoustic spectroscopy • WFPS, Water-filled pore space

	INTRODUCTION

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
APPLICATION of high amounts of fertilizers and poor quality irrigation water may cause an accumulation of soluble salts in agricultural soils. Soluble salt concentrations (e.g., Ca²⁺, Mg²⁺, K⁺, Na⁺, H⁺, NO₃^–, SO₄^2–, Cl^–, HCO₃^–, CO₃^2–, OH^–) in soils are generally measured in terms of electrical conductivity (EC) (Rhoades, 1993, Smith and Doran, 1996). Most soils are considered slightly saline if the EC of a saturated paste extract exceeds 2 dS m^–1 (Smith and Doran, 1996). Microbial growth, nitrogen transformations, and decomposition of organic matter are affected by high concentrations of dissolved cations and anions in soils (Mendum et al., 1999; Avrahami et al., 2002; Killham, 1985; Irshad et al., 2005; Laura, 1974; Frankenberger and Bingham, 1982).

Microorganisms vary widely in their tolerance to salt stress, but bacterial processes such as nitrification and denitrification can be greatly affected by soil EC at levels that are well below the commonly used salinity thresholds (Smith and Doran, 1996). In the absence of soil salinity, changes in soil nutrient levels influence soil EC through differences in the type and number of cations and anions held by the soil particles. Soil water content influences soil EC through the concentration of dissolved ions in the soil. When soil water content is high, dissolved ions (solutes) are diluted; when soil water content is low, dissolved ions are concentrated. This relationship also affects water and substrate supply to microbial cells. To adapt to increased soluble salts at low soil water content, microorganisms create internal solute concentrations either by producing compatible organic solutes or by taking up ions from the extracellular solution (Csonka, 1989). Since water availability is an important factor influencing microbial cells, microbial processes such as nitrification, denitrification and soil respiration are greatly affected.

Application of fertilizer N often leads to transient, localized, and high concentrations of soil solutes. High salt concentrations may cause an inhibition of nitrification after the addition of fertilizer N. Significant increases in N₂O emissions may particularly occur when fertilizer N is applied in excess of crop N demand. More generally, fertilizer effects on N₂O emission can vary widely, depending on factors such as soil type, the amounts and forms of N applied, or N application methods (Granli and Bockman, 1994; Eichner, 1990). Application of 56 to 224 kg N ha^–1 as NH₄NO₃ to barley plots in northeastern Colorado resulted in total N₂O emissions that averaged 0.5% of the fertilizer added (Mosier et al., 1982). In another study, the gaseous N loss associated with the addition of Ca(NO₃)₂ or urea (120 kg ha^–1 N) to a maize field over an 85 d period of sampling was 0.33 and 2.5% of the N applied, respectively (Duxbury and McConnaughey, 1986). Likewise, Breitenbeck et al. (1980) observed that emissions of N₂O-N during a period of 96 d from soil treated with NH₄⁺ fertilizers (125–250 kg N ha^–1) were 0.11 to 0.14% of the N applied as compared to only 0.01 to 0.4% for Ca(NO₃)₂. McSwiney and Robertson (2005) reported that 7% of fertilizer N was lost as N₂O at an N application rate of 134 kg N ha^–1 in a continuous maize cropping system managed at relatively low yield levels. Total N₂O emission increased for N applications greater than 134 kg N ha^–1, but decreased to 2 to 4% of the fertilizer N applied in relative terms. The authors speculated that the drop of N₂O flux was due to a change in the microbial processes that produced N₂O and controlled N availability.

Many field studies suggest that significant soil N₂O fluxes are most likely to occur when water-filled pore space (WFPS) exceeds 60%, soil nitrate concentration is >5 mg NO₃–N kg^–1 dry soil, and soil temperature is >5°C (Abbasi and Adams, 2000; Dobbie and Smith, 2003; Kessavalou et al., 1998; Qian et al., 1997; Sehy et al., 2003; Simojoki and Jaakkola, 2000). At constant soil water content and temperature, rates of soil N₂O and CO₂ production and emission are primarily associated with substrate supply for microbial respiration, nitrification, denitrification, and other external factors that may influence the microbial community. Solute salt concentration, measured as EC, is one of those external factors, but its direct influence on soil greenhouse gas emissions is little understood.

Linear relationships between EC and soil nitrate content in nonsaline soils have been observed in several studies. Smith and Doran (1996) found high positive correlations between EC and NO₃–N for fall and summer measurements in a winter wheat–dry pea rotation. Patriquin et al. (1993) also observed a linear relationship between EC and soil nitrate content in shallow groundwater of a maize field in Canada. The presence of NO₃^– is one prerequisite for N₂O production. Since soil NO₃–N content tends to be positively correlated with N₂O emissions (Mosier et al., 1983), we may assume that a close relationship also exists between soil EC and N₂O emissions.

Although there are many studies of specific ion and soil water effects on microbial processes, the range of soil EC (2.5 to 20 dS m^–1) used was mostly outside EC levels found in agricultural soils (Wijler and Delwiche, 1954; Irshad et al., 2005; Heilman, 1975; Dinesh et al., 1995; Darrah et al., 1987; Frankenberger and Bingham, 1982; McClung and Frankenberger, 1985). No studies that we are aware of have investigated the effect of soil EC < 2.0 dS m^–1 and soil water content at relatively low levels of soil NO₃–N content on greenhouse gas production in soil. One reason may be that it is difficult to separate the role of soil EC from soil NO₃–N content, since these factors are often interdependent. Amos et al. (2005) conducted field measurements of N₂O and CO₂ fluxes in an irrigated maize system. Based on different sampling events during the growing season, they reported a positive correlation between N₂O emission and soil EC at WFPS of 54 to 68%. However, in their case, high N₂O emissions coincided with high fertilizer input, i.e., the increased EC resulted from fertilizer applications and the high N₂O fluxes may have been caused by high soil NO₃–N levels, not necessarily due to a direct effect of EC on microbial activities.

Our hypothesis was that soil N₂O emissions are increased at high soil EC levels by shifting microbial N transformations toward pathways that favor N₂O production and that this occurs in the presence of relatively low initial soil nitrate content. A laboratory experiment was conducted to (i) quantify how soil EC and water content affect the emissions of N₂O and CO₂ in agricultural soils and (ii) determine the ranges of initial soil EC that inhibit or increase N₂O production at low levels of nitrate concentration in the soil.

	MATERIALS AND METHODS

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Soils
Five soils representing different intensive maize-based cropping systems were used in this study (Tables 1 and 2). Two fresh samples of Kennebec silt loam soils (fine-silty, mixed, superactive, mesic Cumulic Hapludolls) (0 to 0.3 m depth) were collected from the Ecological Intensification of Irrigated Maize-based Cropping Systems Experiment located at the University of Nebraska in Lincoln, NE (40°82' N; 96°65' W; elev. 357 m). This experiment was established in 1999 to identify efficient crop management practices for achieving yields that approach yield potential levels and to determine resource-use efficiencies and greenhouse gas fluxes in intensively managed maize and soybean systems (Amos et al., 2005). Samples were collected from two contrasting cropping systems (Table 2): (i) Lin-CC: continuous maize with intensive management (high plant density, intensive NPK management) and (ii) Lin-CS: maize–soybean rotation with recommended best management (normal plant density, recommended NPK management). In each treatment replicate plot, three soil cores (33 mm i.d.) were taken between maize rows after harvest in 2005. The twelve soil cores were thoroughly mixed to form one composite sample per treatment.

The Floyd–Webster loam (Man-CC) soil (fine-loamy, mixed, superactive, Aquic Hapludolls) was collected from the farm of Francis Childs, a frequent winner of maize yield contests, located at Manchester, Iowa (42°47' N; 91°45' W; elev. 302 m). The field is under rainfed continuous maize cropping for >30 yr. Crop management practices include application of 400 to 500 kg N ha^–1 and high plant density (Murrell and Childs, 2000). Soil samples were collected from three 7 by 12 m strips of a high-yield demonstration plot managed by Mr. Childs. Twelve soil cores (33 mm i.d.; 0 to 0.3 m depth) were taken between maize rows at harvest stage of maize in 2002 and thoroughly mixed to form one composite sample.

Laboratory Incubations
Two L of deionized water was added to 1.2 kg of each soil and the soils were placed on a shaker for 30 min to remove soluble ions. After shaking, the soil suspension was allowed to stand for 1 to 2 h. Standing water was removed and the remaining soil paste was tested for soil EC using a pencil conductivity probe (Hanna Instruments Dist WP, Woonsocket, RI). Soil washing was repeated for several times until soil electrical conductivity measured in 1:1 soil/water suspension (EC_1:1) reached 0.5 dS m^–1. The soils were air-dried for at least 3 d to a volumetric water content (_V) of <5%. After drying, the soils were crushed, passed through a 2-mm sieve, and stored in airtight glass containers before soil incubation. Ten grams of soil (dry weight basis) were weighed and 80 mL of 2 M KCl was added. The suspension was shaken for 30 min and filtered through a Whatman no. 42 filter paper. Soil NO₃–N contents in soil extracts were measured using the Cd-reduction method (Mulvaney, 1996), whereas soil NH₄–N contents were determined by the salicylate method (Kempers and Zweers, 1986). Both soil NO₃–N and NH₄–N contents were expressed in mg N per kg dry soil, and the results are summarized in Table 1. A 50-g sample of soil was analyzed for soil texture and soil organic C (Table 1).

Soil cores for incubation were prepared as follows. Fifty grams of each soil (dry weight basis) were weighed into a 100-mL beaker and compacted to a bulk density of between 1.09 and 1.14 Mg m^–3. Each of the soils was treated with KCl, CaCl₂ and Na₂SO₄ salt mixtures dissolved in deionized water at rates that produced electrical conductivities of the bulk soil of approximately 0.5, 1.0, 1.5, and 2.0 dS m^–1 and water-filled pore space of 60 and 90%. All salt treatments had an ionic ratio of Cl^–: SO₄^2–: K⁺: Ca²⁺: Na⁺ = 3: 2: 1.5: 1: 1. The EC values were achieved by carefully pipetting 1 to 8 mL of salt solution per 50 g of soil on a dry weight equivalent basis into a test tube. The salt solution was brought to a final volume by adding deionized water that produced a water-filled pore space of 60 and 90%. The salt solution was shaken using a vortex mixer and gently poured into the soil. Before soil incubation, each soil was tested to estimate the amounts of salt and volume of deionized water that produced 0.5, 1.0, 1.5, and 2.0 dS m^–1 and WFPS of 60 and 90%. Water contents of the five air-dried soils were determined by drying soils for at least 2 d at 105°C. The volume of water added to salt solution per soil core was adjusted using the equation:

[1]

where _g at % WFPS is gravimetric soil water content at 60 or 90% WFPS (%), WFPS is the fraction of pore space filled with water (%), BD is the bulk density of the soil core, and 2.65 is the soil particle density (Mg m^–3). The salt concentrations used for each soil ranged from 0.009 to 0.477 M KCl, CaCl₂, and Na₂SO₄ at 60% WFPS and 0.001 to 0.144 M KCl, CaCl₂, and Na₂SO₄ at 90% WFPS. We chose K⁺, Ca²⁺, Na⁺, SO₄^2–, and Cl^– ions as electrolytes because these are common ions in many cultivated soils and in irrigation water. Soil water contents of 60 and 90% WFPS were chosen because these levels represent previously reported thresholds for maximum microbial nitrification and denitrification, respectively (Linn and Doran, 1984). The soil cores were placed individually inside a 2-L glass jar with a screw-cap lid in which a septum was fitted for gas sampling. The jars were incubated at 25°C for 10 d. Each treatment was replicated three times in a randomized complete block design. For the 90% WFPS treatment, another duplicate set of 50 g soil cores was repacked and treated with salt solution for measurement of soil redox potential.

Gas and Soil Analyses
During the 10-d incubation, gas samples of the headspace of all jars were analyzed daily for N₂O and CO₂ concentrations using a Model 1312 infrared photoacoustic spectroscopy (PAS) gas analyzer (Innova Air Tech Instruments, Ballerup, DK). All measurements were done using a closed-loop system. The jar headspace was sampled using a flow-through lid consisting of rubber septa on jar lids and a 5-cm stainless infusion needle (gauge # 19) connected to the inlet and outlet ports of the PAS gas analyzer. Since the needles used for gas analyses were impervious to air and water, this setup permitted a closed-loop system in which an equivalent amount of air was returned to the jar through the outlet port of the PAS gas analyzer every time a gas sample was drawn through the inlet port. Immediately after the end of each gas analysis, the jars were flushed with atmospheric air for 3 min before the jars were resealed with silicon sealant and returned to the incubator. Water content of soil cores was maintained by weighing and adding necessary deionized water to incubating soil cores. Control samples were included to correct for small amounts of N₂O and CO₂ contained in jars without soils. The control jars contained atmospheric N₂O of 0.262 to 0.382 mg L^–1 and ambient CO₂ of 482 to 516 mg L^–1. There was no system backpressure every time the PAS gas analyzer drew air from the headspace, suggesting that the 1 d incubation did not cause over-pressurization of the jar. The PAS instrument was set at 1-min intervals and measurements of N₂O and CO₂ concentrations in each jar were conducted in triplicates. The production of N₂O and CO₂ was estimated by subtracting the amount of the N₂O and CO₂ in control jars from the amount of N₂O and CO₂ evolved from soil cores. The results were averaged and expressed in terms of gas concentration. The N₂O and CO₂ emission rates were calculated as follows:

[2]

where F is the gas production rate for N₂O (g N₂O-N m^–2 d^–1) or CO₂ (g CO₂–C m^–2 d^–1), C/t denotes the change in gas concentration in the headspace (10^–6 L L^–1 d^–1), V is the headspace volume of the jar (L), A is the surface area of soil core (m²), is the density of gas at 20°C and 0.101 M Pa (1 mol per 24.04 L), and is a conversion coefficient (28/44 for N₂O and 12/44 for CO₂).

Redox potentials of the soil cores were measured after 1-, 3-, 7-, and 10-d incubation periods using an Eh meter (Orion 525A+ meter, Thermo Electron, USA). A platinum electrode was placed at the middle of the soil core for at least 30 min or until a relatively stable readout was obtained. The Eh measurements were expressed in mV and are presented in Table 3.

Statistical Analyses
The five soils, four levels of soil EC and two water contents were replicated three times in a 5 x 4 x 2 x 3 factorial design. Statistical analyses were performed using the General Linear Model procedure (proc GLM) in SAS (SAS, 1999). An analysis of variance (ANOVA) was performed to determine significant differences of N₂O and CO₂ production at different levels of soil EC and water content among all soils and within each soil. Least significant difference values (LSD) were calculated at P < 0.05, when the effects of soil EC and WFPS on N₂O and CO₂ production were found to be significant. All analyses were performed using proc GLM (SAS, 1999).

	RESULTS

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Mineral Soil Nitrogen Content and Soil Electrical Conductivity
All soils had low levels of NO₃–N before salt additions. After a series of soil washings, initial mineral soil content ranged from 0.26 to 1.76 mg NO₃–N kg^–1 and 10.3 to 26.9mg NH₄–N kg^–1 (Table 1). Incubation at 60% WFPS resulted in an increase in NO₃–N compared to initial levels, but a negative relationship between final NO₃–N content and soil EC (Table 3). Final NO₃–N was lowest at EC levels greater than 1.5 dS m^–1, particularly in the Mead and Manchester soils (P = 0.006). At 90% WFPS, final NO₃–N levels in all soils were very low and not affected by EC, indicating nearly complete loss of nitrate by denitrification (Table 3). On average, soil EC after 10 d incubation at 60 or 90% WFPS decreased only slightly (<10 to 15%) from the initial EC levels that had been established by salt addition (Table 4).

View larger version (18K): [in this window] [in a new window]   Fig. 1. Ratio of soil NO3–N and NH4–N contents in all soils at different levels of soil EC and water contents after a 10-d incubation at 25°C.

View larger version (35K): [in this window] [in a new window]   Fig. 2. Cumulative N2O emissions in all soils at 60% water-filled pore space (WFPS) and four levels of soil electrical conductivity (EC). Vertical bars show standard errors. Values in parenthesis represent the final soil EC (dS m–1) measured after a 10-d incubation at 25°C. Letters at the end of each line indicate significance of differences by LSD test at = 0.05.

View larger version (38K): [in this window] [in a new window]   Fig. 3. Measured daily N2O and CO2 flux rates in Lin-CC soil at 60 and 90% water-filled pore space (WFPS) and four levels of soil electrical conductivity (EC). Vertical bars show standard errors. Values inside parenthesis show the final soil EC (dS m–1) measured after a 10-d incubation at 25°C.

View larger version (32K): [in this window] [in a new window]   Fig. 4. Cumulative CO2 emissions in all soils at 60% water-filled pore space (WFPS) and four levels of soil electrical conductivity (EC). Vertical bars show standard errors. Values in parenthesis represent the final soil EC (dS m–1) measured after a 10-d incubation at 25°C. Letters at the end of each line indicate significance of differences by LSD test at = 0.05.

View larger version (36K): [in this window] [in a new window]   Fig. 5. Cumulative N2O emissions in all soils at 90% water-filled pore space (WFPS) and four levels of soil electrical conductivity (EC). Vertical bars show standard errors. Values in parenthesis represent the final soil EC (dS m–1) measured after a 10-d incubation at 25°C. Letters at the end of each line indicate significance of differences by LSD test at = 0.05.

Average cumulative CO₂ production was similar to that measured at 60% WFPS and it also decreased from a high of 4.4 g CO₂–C m^–2 at 0.5 dS m^–2 soil EC to a low of 3.1 g CO₂–C m^–2 when soil EC was raised to 2.0 dS m^–2 (Table 4, Fig. 6). Daily CO₂ flux rates increased after 1 d and mostly peaked around 4 d of incubation (Fig. 4). There was a decrease in the respiration rate after an 8 d incubation in all treatments and soils, presumably due to exhaustion of C substrates. Similar to 60% WFPS, highest CO₂ losses were measured in the Man-CC soil.

View larger version (32K): [in this window] [in a new window]   Fig. 6. Cumulative CO2 emissions in all soils at 90% water-filled pore space (WFPS) and four levels of soil electrical conductivity (EC). Vertical bars show standard errors. Values in parenthesis represent the final soil EC (dS m–1) measured after a 10-d incubation at 25°C. Letters at the end of each line indicate significance of differences by LSD test at = 0.05.

	DISCUSSION

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

At 60% WFPS, the short-term decrease in N₂O flux after a 1-d incubation in all soil EC treatments and soils suggests that low initial soil NO₃–N and low nitrification rates limited microbial N₂O production. As more nitrate was produced over time by nitrification, N₂O flux rates started to increase after a 3-d incubation. The increase in daily respiration rate after a 1-d incubation also supports the assumption that mineralization of organic matter followed by nitrification was the major process that led to production of N₂O at 60% WFPS. At 90% WFPS, N₂O production is primarily related to changes over time in the rates of nitrification, denitrification, NO₃, and carbon substrate supply. Similar to our incubation study, increased N₂O emissions at high WFPS have been observed in many field studies (Abbasi and Adams, 2000; Qian et al., 1997; Sehy et al., 2003; Simojoki and Jaakkola, 2000). Although the measured soil redox potential indicated that at 90% WFPS conditions for denitrification started to prevail in bulk soil within a few days (Table 3), there were probably sufficient microsites to sustain simultaneously high rates of both nitrification and denitrification, particularly near the soil surface where O₂ concentration was high, resulting in high flux rates during the first days of incubation. The variation among soils in the patterns of N₂O emission at 90% WFPS (Fig. 5) was probably caused by differences in such microsites and in substrate supply among the experimental soils, including the differences in the NO₃ supply relative to the available C substrate (Weier et al., 1993).

Our study showed that the N₂O and CO₂ production by the microbial community were affected by salt concentration (in situ soil EC) at both water contents and even at low soil nitrate content. Increasing soil EC decreased heterotrophic soil respiration at 60 and 90% WFPS, suggesting a general reduction in microbial activity due to increased osmotic stress on the microbial communities. Killham (1985) also found a decrease in respired C and dehydrogenase activity when a sandy loam soil was irrigated with increasing salinity. Frankenberger and Bingham (1982) reported that dehydrogenase activity was severely inhibited (30 to 81%) when the EC of a soil extract (EC_e) was increased from 0.2 to 22 dS m^–1. Soil CO₂ production at various EC levels was positively correlated with soil organic C content (P < 0.01), which has also been found in other studies (Zaman et al., 2004).

The observed interactions between increasing salt concentrations, water content, and N₂O emissions from soils are likely to be related to the physiological status of microbes involved in nitrification and denitrification. Largest NH₄–N concentration was generally observed at 60% WFPS and an EC of 2.0 dS m^–1, whereas it was lowest in soils with low EC (Table 3). This suggests that, unlike C mineralization (Fig. 4), N mineralization was not suppressed by increasing EC, whereas nitrification of ammonium to nitrate was inhibited by increasing salt concentrations. Similar observations have been made in a number of studies with widely varying EC levels, including saline and alkaline soils (Laura, 1974; McClung and Frankenberger, 1985; McCormick and Wolf, 1980; Pathak and Rao, 1998). Added salt mixtures increase the concentration of ions in the soil solution and decrease available water for microbial activity. To survive under stress, soil microbial communities change their microbial functions and/or activity in response to environmental disturbances. Several reports suggest that bacteria involved in nitrification regulate intracellular solute concentrations through accumulation and loss of amino acids (Schall et al., 1981; Killham and Firestone, 1984). More generally, the ability to do this may vary among microbial communities. Studies by Stark and Firestone (1995) indicated that there was a 75% decrease in nitrification rates when soil was exposed to 0.67 M K₂SO₄ solutions. Microbial nitrification was affected because the bacteria involved were unable to produce compatible solutes as the cell underwent dehydration. In our study, this would explain the general reduction in N₂O emissions with increasing EC at 60% WFPS. Other indications for this include the lack of significant NO₃–N accumulation at EC > 1 dS m^–1 (Table 3) and slightly decreasing final soil EC values (Table 4).

Microbial cells may also employ another response mechanism to salt stress by catabolizing intracellular solutes to CO₂ or polymerizing them into osmotically less active compounds (Halverson et al., 2000; Avron and Ben-Amotz, 1979; Reed and Stewart, 1983). Considering this, it is possible that denitrifiers may employ several response mechanisms to salt stress that lead to a generally greater tolerance than that of nitrifiers. Halverson et al. (2000) reported that in response to addition of 0.11 M NaCl, Pseudomonas spp. released a maximum of 22 to 26% of their amino acid pool and 11 to 21% of their low molecular weight in neutral sugars. Korber et al. (1996) reported that Pseudomonas fluorescens, a common denitrifying bacteria in soils, plasmolyzed on the addition of 1.0 to 1.5 M NaCl, but were able to resume growth after removal of the salt stress. In our study, it appears that at 2.0 dS m^–1 initial soil EC and 90% WFPS, where the salt concentration was a 0.03 to 0.1 M KCl, Na₂SO₃, and CaCl₂ mixture, bacterial communities dominated by denitrifiers performed their metabolic functions better than nitrifying bacteria at 60% WFPS by diluting the external salt stress through response mechanisms such as those discussed above. Another indirect indication for a more efficient response of denitrifiers to higher salt concentrations is the somewhat greater decline in EC and the overall large loss of soil NO₃ observed in all soils during the 10-d incubation at 90% WFPS.

With the salt additions used in this study, the presence of specific ions may have also caused nutritional imbalances that affected microbial growth rates and enzyme activities. Several studies noted that reduction of nitrification rates was more severe in the presence of chloride salts when compared to the sulfates added at equivalent rates (Frankenberger and Bingham, 1982; Dinesh et al., 1995; McClung and Frankenberger, 1985; Agarwal et al., 1971). They attributed the lower degree of inhibition by sulfates to the enhanced utilization of sulfate anions in microbial cells for the synthesis of amino acids, vitamins, sulfolipids, and several other essential organic compounds. In contrast, chloride ions are less likely to be incorporated in anabolic pathways and hence may accumulate in soils at toxic levels. In our study, because Cl^– and SO₄^2– ions were used in the salt mixtures and the concentration of Cl^– ions in each EC levels was relatively high, toxic effects of chloride on soil microbes may have contributed to reduced N₂O production when salt concentration increased from 0.01 (0.5 dS m^–1) to 0.48 (2.0 dS m^–1) M salt mixtures at 60% WFPS. However, at high water content (90% WFPS) significant nitrate production and denitrification occurred at comparable levels of salt addition, indicating that inhibition of nitrification by Cl^– and SO₄^2– ions was probably negligible. The data presented here do not allow a more detailed interpretation of potential selective toxicities of specific ions.

The range of initial bulk soil EC chosen for this study, 0.5 to 2.0 dS m^–1, was relatively narrow, but it represents conditions found in many agricultural soils. It is also typical for temporary or localized increases in EC that may result from fertilizer application (Amos et al., 2005). Within that range, effects of EC on N₂O and CO₂ emissions were most significant at initial EC levels of >1 dS m^–1 and in soils with greater biological activity. Our experiment did not allow for the derivation of robust quantitative relationships between CO₂ or N₂O fluxes and soil EC that would be applicable to field conditions, including the presence of growing plants. More research should be conducted on this topic so that direct effects of soil EC on soil greenhouse gas production can also be accounted for in ecosystem simulation models that aim to predict greenhouse gas emissions from soil. Because we did not measure community structures of ammonia oxidizers and denitrifiers or the physiological or metabolic state of the microorganisms at increasing soil EC levels, the above discussion on mechanisms of soil EC effects on trace gas evolution at different levels of water content requires further verification in the field. In particular, the mechanisms used by soil microorganisms in response to stress (soil EC > 1.0 dS m^–1) and the types of microorganisms proliferating during high salt conditions are not well understood.

	CONCLUSIONS

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
In the presence of low soil NO₃–N content, soil EC affects microbially driven production of greenhouse gases, but its direct influence varies according to water content and EC level. Average microbial respiration decreased with increasing the initial soil EC from 0.5 to 2.0 dS m^–1 regardless of soil water status. At 60% WFPS, N₂O production decreased with increasing EC in all soils. At 90% WFPS, N₂O production was greater than that at 60% WFPS and N₂O losses increased with increasing soil EC. Differences in greenhouse gas emissions due to varying soil EC and WFPS may be related to changes in mechanisms of adjustment to salt stress by microbial communities and microbial toxicities to specific ions. The general decrease in soil respiration with increasing soil EC may have been due to osmotic stress on microbial activities. At 60% WFPS, the reduced N₂O production at increased EC was most likely associated with decreasing activity of nitrifying bacteria, which became stressed at higher EC levels. However, when soil water content was raised to 90% WFPS, denitrification was the most likely primary source of N₂O and the microbial community involved in the process appears to be more tolerant to salt stress. The major implication arising from our laboratory study is that ecosystem models that attempt to simulate N₂O and CO₂ fluxes from soils may have to also account for general or short-term effects of rising or declining soil EC on microbial activities. Whether the relationships shown in the laboratory are of equal importance under field conditions remains to be verified. Likewise, better understanding of physiological shifts in the soil microbial community in response to increasing soil EC or water content is needed.

	ACKNOWLEDGMENTS

 
We thank Spencer Arnold and Susan Siragusa for providing soils and instruments used in this study and two anonymous reviewers for their helpful suggestions. This research was supported by funds provided by U.S. Department of Energy: a) EPSCoR program, Grant No. DE-FG-02-00ER45827 and b) Office of Science, Biological and Environmental Research Program (BER), Grant No. DE-FG03-00ER62996.

	NOTES

TOP NOTES ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION CONCLUSIONS REFERENCES

 
Contribution of the Nebraska Agric. Exp. Stn. Scientific J. Series Paper no. 15188.

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