Published online 28 June 2005
Published in Soil Sci Soc Am J 69:1233-1237 (2005)
DOI: 10.2136/sssaj2004.0283
© 2005 Soil Science Society of America
677 S. Segoe Rd., Madison, WI 53711 USA
Soil Biology & Biochemistry Note
ATMOSPHERIC NITRATE DEPOSITION AND ENHANCED DISSOLVED ORGANIC CARBON LEACHING
TEST OF A POTENTIAL MECHANISM
Jared L. DeForesta,b,*,
Donald R. Zaka,c,
Kurt S. Pregitzerd and
Andrew J. Burtond
a School of Natural Resources & Environment, Univ. of Michigan, Ann Arbor, MI 48109-1115
b Current address: Earth, Ecological and Environmental Sciences, Univ. of Toledo, Toledo, OH 43606-3390
c Dep. of Ecology and Evolutionary Biology, Univ. of Michigan, Ann Arbor, MI 48109-1048
d School of Forest Resources and Environmental Science, Michigan Technological Univ., Houghton, MI 49931-1295
* Corresponding author (Jared.DeForest{at}utoledo.edu)
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ABSTRACT
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Atmospheric NO3– deposition has the potential to disrupt litter decomposition in temperate forests by suppressing enzymes responsible for lignin degradation. A reduction in phenol oxidase activity could potentially trigger an increase in soluble phenolic compounds, which in turn are known to decrease the activity of cellulolytic enzymes like ß-glucosidase. Our study investigated whether the inhibition of lignin degradation by experimental NO3– deposition could increase soluble phenolics in soil, suppress ß-glucosidase activity, and potentially explain a greater export of dissolved organic C (DOC) from northern hardwood ecosystems. We found no evidence that the suppression of phenol oxidase by NO3– additions increased soluble phenolics in mineral soil, nor did we find a strong inverse relationship between soluble phenolics and ß-glucosidase activity. It appears that reductions in mineral soil lignolytic activity induced by experimental deposition are not responsible for greater DOC export from soil.
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INTRODUCTION
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CHRONIC ATMOSPHERIC NO3– deposition has the potential to directly affect microbial activity and initiate a series of physiological changes that alter the cycling and storage of C in soil (Berg, 1986; Berg and Matzner, 1997). For example, white rot Basidiomycota and xylariaceous Ascomycota are the primary agents of lignin degradation, and high concentrations of inorganic N can inhibit their ability to produce phenol oxidase and peroxidase (Keyser et al., 1978; Kirk and Farrell, 1987; Fog, 1988). Experimental atmospheric NO3– additions in forest ecosystems, at levels already occurring in some portions of the northeastern USA and Europe (Fenn et al., 1998), have caused significant reductions in the activity of these extracellular enzymes (Carreiro et al., 2000; Saiya-Cork et al., 2002; DeForest et al., 2004a). Recently, we have observed that experimental NO3– deposition also can reduce ß-glucosidase activity (DeForest et al., 2004a) and increase the leaching of DOC from soil (Pregitzer et al., 2004), both of which occurred with a concomitant decline in lignolytic activity. These observations suggest that chronic atmospheric NO3– deposition can set in motion a series of microbial physiological changes that diminish the degradation of lignified plant cell walls and increase microbial byproducts of plant litter degradation (e.g., DOC). An increase in the DOC leaching would likely increase the leaching of soluble phenolics, which is a component of the total DOC pool. Here, we test a potential mechanism by which chronic NO3– deposition could elicit this chain of events.
Our rationale linking declines in extracellular enzyme activity to increases in soluble phenolic leaching centers on NO3– deposition reducing the activity of lignolytic enzymes, resulting in the partial degradation of lignin in litter and humified compounds in soil organic matter. Such a response would be consistent with our previous observations (DeForest et al., 2004a), and it could lead to the accumulation of soluble phenolics in soil solution, which are known to inhibit the activity of extracellular enzymes responsible for cellulose hydrolysis (i.e., ß-glucosidase). In wetland soils, low pO2 limits phenol oxidase activity (McLatchey and Reddy, 1998), which can produce an accumulation of soluble phenolics and the suppression of ß-glucosidase and other cellulolytic enzymes (Freeman et al., 2001). Copolymerization and immobilization into soil organic matter are two processes by which soluble phenolics and polyphenolics (e.g., tannins) might inhibit ß-glucosidase and other hydrolytic enzymes (Sarkar and Burns, 1983). We hypothesized that a similar series of events is initiated by atmospheric NO3– deposition in upland forest soils, in which phenol oxidase and peroxidase activity have been suppressed. Greater production of soluble phenolics by NO3– deposition also might explain substantial increases in DOC leaching that we have previously observed (Pregitzer et al., 2004).
To test this mechanism, we measured phenol oxidase, peroxidase, ß-glucosidase, and soluble phenolic concentrations in the mineral soil of northern hardwood forest sites receiving ambient and experimental atmospheric NO3– deposition. If this mechanism was at work in our experiment, we expected to observe an inverse relationship between the activity of lignolytic enzymes and soil phenolic concentrations, as well as an inverse relationship between phenolic concentrations and ß-glucosidase. We expected experimental NO3– deposition would produce significantly higher concentrations of soluble phenolics and lower activity of extracellular enzymes involved with lignin and cellulose degradation.
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Materials and Methods
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Study Area and Field Sampling
To determine soil phenolic concentrations and their potential to suppress ß-glucosidase activity, we studied soil in four Acer saccharum Marsh.—dominated northern hardwood sites in Michigan with similar stand composition, history, structure, and soil development (Fig. 1
; Burton et al., 1991). Soils at the three northern sites (A, B, and C) were sandy, mixed, frigid Typic Haplorthods and sandy, mixed, frigid Alfic Haplorthods. At site D, soils were sandy, mixed, mesic Entic Haplorthods. These study sites are located along a 500-km climatic and N deposition gradient where the northernmost Site A receives the least amount of N deposition (6.8 kg N ha–1 yr–1), followed by Site B at (9.1 kg N ha–1 yr–1), Site C and Site D around (11.7 kg N ha–1 yr–1). These sites represent a common forest ecosystem found throughout the Upper Great Lakes region. Within each study site, six 30 m x 30 m experimental plots were established. Three plots receive ambient levels of N deposition and served as a control. Since 1994, the other three plots received 30 kg NO3––N ha–1 yr–1 in addition to ambient N deposition. We broadcast six equal applications of dry NaNO3 over the forest floor during the growing season to simulate atmospheric NO3– deposition. The amount and type of N added is similar to atmospheric deposition in forests near industrial regions of the northeastern USA (Fenn et al., 1998).
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Fig. 1. Distribution of the four northern hardwood sites along a 500-km climatic and NO3– deposition gradient in Michigan, USA. These stands span the geographic distribution of sugar maple dominated northern hardwood forests in the upper Great Lake States region.
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We sampled the soil six times during the 2002 growing season (May to October) to assess the influence of experimental NO3– deposition on soluble soil phenolics and extracellular enzyme activity. On each sampling date, we collected eight soils cores (2-cm diam. and 10 cm deep) in plots receiving ambient and elevated atmospheric NO3– deposition. The surface organic horizons (Oi and Oe) were removed and the soil cores contained a small amount of the Oa horizon (<0.5 cm) and surface mineral soil (0–10 cm) of A and E horizon. The eight soil samples from each plot were composited, and all subsequent analyses were performed on the composite samples. Samples were kept on ice and processed within 48 h of field collection.
Extracellular Enzyme Activity
We used three subsamples of each composite sample to assay the activity of phenol oxidase, peroxidase, and ß-glucosidase. The enzyme assays were prepared by mixing 2 g of soil in 150 mL of acetate buffer (50 mM, pH 5). ß-glucosidase activity was fluorometrically measured in 96-well plates using 4-methylumbelliferone-ß-D-glucoside as a substrate (Saiya-Cork et al., 2002). At the termination of the assay, we added 25 µL of NaOH (0.2 M), to each well and then measured fluorescence using a F-max fluorometer (Molecular Devices Corp., Sunnyvale, CA); excitation energy was 355 nm and emission was measured at 460 nm. We measured peroxidase and phenol oxidase activity using L-dihydroxyphenylalanine (Saiya-Cork et al., 2002). After incubating the samples at 25°C for 18 h in 96-well plates, the optical density (460 nm) of the oxidized reaction product was measured on a spectrophotometer (Bio-Tek Instruments, Winooski, VT). All enzyme activities are expressed as nanomoles of substrate cleaved per gram dry mass of soil per hour (nmol g–1 soil h–1).
Soluble Soil Phenolics
Soluble soil phenolic concentrations were determined by comparing soil solution with a standard mixture of phenolic compounds. This standard mixture contained 50 µmol L–1 each of ferulic, p-coumaric, p-hydroxybenoic, vanillic, and syringic acid. We adjusted the standard mixture to pH 6 and then diluted the standard to have a graduated range of phenolic concentrations from 3 to 250 µmol L–1, which encompasses a common range of phenolic compounds and concentrations in soil (Sposito, 1989). To extract the soluble phenolics from soil, we agitated 5 g of soil and 25 mL of water on an orbital-action shaker for 18 h. The samples were centrifuged (900 x g) and the supernatant was passed through a 0.45-µm nylon filter (Ohno and First, 1998). We combined 5 mL of filtered supernatant, or standard phenolic mixture, with 0.75 mL of Na2CO3 (1.9 M) and 0.25 mL of Folin-Ciocalteu reagent (Ohno and First, 1998). Deionized water was used as a negative control. After 1-h incubation at 25°C in the dark, absorbance was measured at 750 nm using a Spectronic 20 Genesis spectrophotometer (Spectronic Instruments, Rochester, NY). A regression of absorbance and standard phenolic concentration was determined (r2 = 0.99), and the absorbance of samples was adjusted to represent phenolic concentrations (µmol C g–1 of soil).
Statistical Analyses
In a previous study, we found that soil water content had a significant influence on enzyme activity (J. DeForest, unpublished data, 2003), and we used it in this analysis to adjust for any differences in field soil water contents among plots and stands. All values presented for enzyme activities are least square means adjusted for differences in soil water content. We used a two-way repeated measures ANOVA, with soil water as a covariate, to investigate the influence of sample date, site and NO3– deposition treatment on soluble phenolics concentrations in mineral soil. We also explored the relationship between enzyme activity and phenolic concentrations using linear regression analysis. Significance for all statistical analysis was accepted at 
= 0.05.
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Results
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Experimental NO3– deposition significantly decreased the activity of phenol oxidase for the two northern sites (A and B) when averaged across sampling dates (Table 1; Fig. 2)
. Although experimental NO3– deposition reduced the activity of ß-glucosidase and peroxidase, these differences were not statistically significant (Table 1); this contrasts with our previous observations of significant declines in activity due to experimental NO3– deposition (DeForest et al., 2004a). Sampling date had a significant (p < 0.001) influence on ß-glucosidase and peroxidase activity, with the greatest enzyme activity occurring in late June. Study sites also were a significant (p < 0.01) influence on enzyme activity (Fig. 2).
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Table 1. Repeated measures analysis of covariance for ß-glucosidase, peroxidase, and phenol oxidase activity; soil water content was the covariate in this analysis. Soluble phenolic concentrations were analyzed using a repeated measures analysis of variance.
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Fig. 2. The influence of experimental NO3– deposition and site on the activity of (A) phenol oxidase and (B) peroxidase, (C) the concentration of soluble phenolics, and (D) the activity of ß-glucosidase. Enzyme activities are least square means, which are adjusted for differences in soil water content. Asterisk represents significance (p > 0.05) and error bars represent standard error of the mean (n = 18).
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Over the growing season, we observed similar concentrations of soluble phenolics in ambient and elevated NO3– deposition treatments (Fig. 3)
, which produce a nonsignificant interaction between these two factors. As a main effect, NO3– deposition had no influence on soluble phenolics (Table 1), with equivalent concentrations occurring under ambient and elevated NO3– deposition for Sites B, C, and D (Fig. 2). Elevated NO3– deposition significantly (p > 0.001) reduced soluble phenolics by 35% at the most northern site (A; Fig. 2). Sampling date and study site had significant influences on phenolic concentrations (Table 1). Soluble phenolic concentrations were generally highest in the spring and fall and lowest in the summer. For example, concentrations peaked in June at 121.5 ± 9.6 µmol C g–1 (mean ± SE) and decreased throughout the growing season to a minimum in August (86.2 ± 5.9 µmol C g–1); by September, phenolic concentrations increased to 111.6 ± 8.3 µmol C g–1. Additionally, phenolic concentrations were highest at the two northern Sites A & B (approximately 126.8 µmol C g–1) and lowest at Sites C and D (approximately 82.3 µmol C g–1). We found a significant negative relationship between phenol oxidase activity and soluble phenolics, but phenol oxidase activity accounted for a small proportion of the variation in soluble phenolic across sites and sampling dates (n = 144; p < 0.001; r2 = 0.09). We also found a significant, negative relationship between soluble phenolics and ß-glucosidase activity, but this relationship also was very weak (n = 144; p < 0.001; r2 = 0.18)
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Fig. 3. Soluble phenolic concentrations in surface mineral soil (y axis) of northern hardwood sites receiving experimental atmospheric NO3– deposition. Although sampling date (x axis) was a significant effect, we found no interaction between NO3– deposition treatment and sampling date. Error bars represent standard error of the mean (n = 12).
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Discussion
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Experimental NO3– deposition produced a significant reduction in phenol oxidase activity; however, we found no effect of NO3– deposition on soluble soil phenolics, indicating that declines in phenol oxidase likely did not produce higher concentrations of these compounds. The fact that soluble phenolic concentrations displayed a very weak inverse relationship with phenol oxidase activity provides further support for this contention. Experimental NO3– deposition produced a nonsignificant decline in ß-glucosidase activity and the activities of this enzyme also displayed a weak inverse relationship with soluble phenolic concentrations in soil. Thus, it appears that reductions in lignolytic activity due to NO3– deposition are unlikely to cause increased soluble phenolic concentration in mineral soil, which could potentially reduce ß-glucosidase activity. Although previous research has demonstrated that high concentrations of phenolics (approximately 2000 µg L–1) can significantly suppress ß-glucosidase activity in poorly drained soils (Freeman et al., 2001), it is possible that the modest concentrations (approximately 800 µg L–1) of soluble phenolics in our fertilized upland soils were insufficient to elicit that same effect. Overall, it appears that ß-glucosidase activity, or other cellulolytic enzymes, is not substantially influenced by soluble phenolics in upland soils receiving NO3– deposition. What alternative mechanisms might be responsible for consistent decreases in extracellular enzyme activity and increases in DOC production in soils receiving experimental NO3– deposition?
The interactions among NO3– deposition, extracellular enzyme activity and the production of DOC are not well understood, but there are several alternatives that may explain the relatively consistent responses we have observed in our previous work. Cellulose is an important substrate for heterotrophic metabolism in soil (Eriksson and Wood, 1985) and a reduction in ß-glucosidase activity could reduce the amount of energy enzymatically derived from cellulose (Desphande et al., 1978). It is plausible that declines in lignolytic capacity limit microbial access to cellulose that is protected by lignin, thus producing a decline in cellulolytic capacity as we have previously observed in mineral soil (DeForest et al., 2004a). This is consistent with the high lignin concentration of sugar maple fine roots (25–36%; Parsons et al., 2003), which represent a substantial input of substrate for microbial metabolism in our study (Hendrick and Pregitzer, 1993). A decline in oxidative capacity would also limit the access of carbohydrates that have been incorporated into soil organic matter, thus also lowering their access to microbial metabolism. These specific responses would be difficult to discern, but they clearly would be accompanied by an increase in soil organic matter, which we have documented in our experiment (DeForest et al., 2004b).
It is unlikely that lower phenol oxidase activity in mineral soil was directly responsible for a three-fold increase in DOC export produced by our NO3– deposition treatment (Pregitzer et al., 2004). Although soluble phenolics represent a fraction of the DOC leaching from this ecosystem, we found no difference in mineral soil soluble phenolic concentrations between ambient and NO3– deposition treatments. Moreover, we observed a very weak inverse relationship between phenol oxidase activity and soluble phenolic concentrations in mineral soil, further suggesting that low rates of lignin oxidation under NO3– deposition do not lead to the accumulation of soluble phenolics in mineral soil. If NO3– induced changes in mineral soil microbial metabolism are not responsible for greater DOC leaching, then what are the mechanisms responsible for this response? The majority of DOC export occurs in late fall and early spring, when fresh leaf litter carpets the forest floor (Pregitzer et al., 2004). The leaves of sugar maple contain substantial amounts of soluble phenolics (Lindroth et al., 1993), and we have observed substantially larger decreases in phenol oxidase (–35%) and peroxidase (–32%) in forest floor, relative to those occurring in mineral soil (–15 and 5%, respectively) in NO3– amended soil; DeForest et al., 2004a). Moreover, the seasonality of leaf litter fall and DOC export also coincides with temporal patterns in soluble phenolics concentrations (Fig. 2). Given the high concentrations of phenolics in sugar maple leaves (Lindroth et al., 1993) and large decreases in oxidative enzyme activity in leaf litter (DeForest et al., 2004a), microbial activity in forest floor may hold the key to understanding why NO3– deposition has increased DOC production and export. It is clear that high concentrations of soluble phenolics are not accumulating in the mineral soil of our NO3– deposition treatment, relative to those occurring in the mineral soil of our ambient treatment.
In conclusion, decreases in phenol oxidase activity did not result in higher concentrations of soluble soil phenolics in mineral soil receiving experimental atmospheric NO3– deposition. Soluble phenolics also displayed a weak negative relationship with ß-glucosidase activity, suggesting that soluble phenolics exert minimal influence the activity of this cellulolytic enzyme. Moreover, a relatively low concentration of soluble phenolics in mineral soil suggests that the production of these compounds is not likely the source of DOC leaching from our NO3– deposition treatment. Although we did not measure forest floor (Oi & Oe) in this study, changes in enzyme activity and DOC production in organic horizons may hold the key to understanding why NO3– deposition has resulted in threefold increase in DOC leaching from this ecosystem. In summary, NO3– deposition did not alter phenolic concentrations in mineral soil, nor were phenolic concentrations strongly related to lignolytic or cellulolytic extracellular enzyme activity. Thus, we have no evidence to support our initial hypothesis that reductions in phenol oxidase due to NO3– deposition would increase soluble phenolics in soil solution, which in turn would inhibit ß-glucosidase and stimulate DOC leaching.
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ACKNOWLEDGMENTS
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This research was supported by the National Science Foundation grants DEB 0315138 and DEB 0075397. Thanks to Matt Tomlinson for site maintenance, Michelle Martin for assistance in the lab, and Jennifer DeForest for field assistance. We acknowledge the USDA Forest Service and the Michigan DNR for providing study site locations.
Received for publication August 24, 2004.
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ABSTRACT
INTRODUCTION
