Soil Biology & Biochemistry 37 (2005) 1469–1475 www.elsevier.com/locate/soilbio
Seasonal protein dynamics in Alaskan arctic tundra soils Michael N. Weintrauba,*, Joshua P. Schimelb,1 a Department of Ecology and Evolutionary Biology, University of Colorado, 334 UCB, Boulder, CO 80309, USA Department of Ecology, Evolution, and Marine Biology, University of California, Santa Barbara, CA 93106, USA
b
Received 6 March 2004; received in revised form 21 December 2004; accepted 4 January 2005
Abstract In the arctic tundra of Alaska, plant growth is limited by N supply, especially in tussock tundra. Because proteins are the predominant form of soil organic N, proteolysis is considered to be the rate-limiting step in both the release of amino acids and in N mineralization. To help understand the controls on N availability in tundra soils, and to determine whether proteolysis is controlled by enzyme activity or by substrate availability, we measured soil protein concentrations, and proteolysis rates with and without added protein, every 1–2 weeks through the summer of 2000 and twice in the summer of 2001. Protease activity with added protein (‘potential’) was higher than without added protein (‘actual’). However, differences between the two tended to be driven by relatively brief peaks in potential protease activity. In fact, actual and potential rates were usually similar, suggesting that much of the time proteolysis was not limited by substrate availability, but rather by enzyme activity. Our data suggest that protease activity was actually only substrate limited at the times when it was highest. Potential rates peaked at the same times that soluble proteins were also high. These increases in protease activity and soluble protein concentrations occurred when soil amino acid and NHC 4 concentrations were at their lowest, drawn down by the seasonal peaks in root growth. The fact that the peaks in protease activity coincided with the peak in soil amino acid and NHC 4 demand strongly suggests that proteolysis was stimulated by high soil amino acid demand, and resulted in increases in soluble protein concentrations caused by the solubilization of larger proteins. q 2005 Elsevier Ltd. All rights reserved. Keywords: Soil proteins; Dissolved organic nitrogen; Tundra soils; Arctic soils; Tundra nitrogen availability; Soil protease activity
1. Introduction In the arctic tundra of Alaska, plant growth is limited by N availability, especially in the tussock community (reviewed in Schimel et al., 1996). Even where soil N content is relatively high, N availability is low because cold temperatures and waterlogged conditions slow the decomposition of organic matter (OM), reducing N release into plant-available forms. Because decomposition rates have been slower than plant production rates (Oechel and Billings, 1992), large amounts of partially decomposed and nutrient poor plant detritus accumulate. Laboratory incubations demonstrate, however, that this stored OM can
* Corresponding author. Tel.: C1 303 492 6248; fax: C1 240 358 6080. E-mail addresses:
[email protected] (M.N. Weintraub),
[email protected] (J.P. Schimel). 1 Tel.: C1 805 893 7688. 0038-0717/$ - see front matter q 2005 Elsevier Ltd. All rights reserved. doi:10.1016/j.soilbio.2005.01.005
decompose rapidly and mineralize significant quantities of N when conditions are favorable for decomposition (Weintraub and Schimel, 2003). It has been argued that increasing temperatures caused by climate warming (Serreze et al., 2000) will increase soil organic matter (SOM) decomposition and N availability, and enhance plant growth (Shaver and Chapin, 1986; Shaver et al., 1992). Because plant material has a higher C-to-N ratio than SOM, and higher N availability will increase plant productivity (Shaver and Chapin, 1980; Shaver et al., 1986), the transfer of N from SOM to plants with warming will make the tundra a stronger sink for atmospheric C, potentially offsetting C losses from SOM decomposition (Rastetter et al., 1992; Shaver et al., 1992). As a result, N supply to plants is one of the primary factors regulating the C balance of the Arctic tundra (Shaver et al., 1992). Net N mineralization has been considered to be the principal control on plant N availability (Vitousek et al.,
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1979; Schimel and Bennet, 2004), however, in the soils of the Alaskan arctic, net mineralization is too low to meet the total N requirements of tundra plants (Giblin et al., 1991; Nadelhoffer et al., 1991). Tundra plants are known to take up amino acids (Chapin et al., 1993; Kielland, 1994; Schimel and Chapin, 1996; Kielland, 1997; Nordin et al., 2004), with amino acid supply rate the critical factor determining amino acid availability to plants (Leadley et al., 1997). Even so, the controls on amino acid production in soil are poorly understood. To understand soil N supply to plants, we need to understand the processes that liberate soil N. Hydrolysable soil organic N consists primarily of insoluble protein-N (Sowden et al., 1977; Nemeth et al., 1988; Schulten and Schnitzer, 1998). Proteins are the largest nitrogenous component of plants (Chapin et al., 1987), and therefore represent one of the largest plant N inputs to soil. They can also be released in soil by microbial turnover (reviewed in Lipson and Nasholm, 2001). However, the majority of soil protein-N is protected by minerals and humus (Lipson and Nasholm, 2001). As a result, the smaller organic N pool of dissolved proteins is likely to be more indicative of the available amino N pool than the largely insoluble pool of total soil protein (Lipson and Nasholm, 2001), and soluble proteins have been found by several researchers to be the principal source of mineralized N (Nemeth et al., 1988; Matsumoto et al., 2000; Finzi and Berthring, 2004). Thus, proteolysis of soil proteins and peptides is considered to be the rate-limiting step in both the release of amino acids and in N mineralization (Ladd and Paul, 1973; Asmar et al., 1994; Lipson et al., 1999b). Proteolytic activity can be controlled by the rate of protein supply to protease enzymes, if they are substrate limited, or by the activity of the enzyme pool, if they are saturated by substrate availability. Schimel and Weintraub (2003) suggested that as the size of the enzyme pool increases, activity per enzyme decreases. As microbes produce more extracellular enzymes that bind to solid substrates, they have to diffuse farther out from the cells, the substrates must diffuse farther back, and the enzymes can compete with one another for binding sites (Reid, 1995; Schimel and Weintraub, 2003). This suggests that enzyme activity is controlled by substrate availability unless the enzyme pool is very small. Thus, we hypothesize that proteolysis in tundra soils is limited by substrate availability. To identify the controls on N availability in different tundra soils over the growing season we determined the relationship between the seasonal patterns of protease activity and protein availability, and whether proteolysis in tundra soils is controlled by enzyme activity or by substrate availability by measuring soluble protein concentrations, and proteolysis rates with and without added protein, every 1–2 weeks through the summer of 2000 and twice in the summer of 2001.
2. Materials and methods 2.1. Study site All sample collection and research work took place at the Toolik Field Station (Lat 688 38 0 N, Long. 149838 0 W), on the north slope of the Brooks Range in Alaska. We sampled three tundra communities: tussock tundra, shrub tundra, and wet sedge tundra. In Alaskan upland tundra, tussock tundra is the most common vegetation type. Tussock is a moist tundra form, which typically occurs on silty to gravelly soils in areas of moderately hilly topography (Shaver and Chapin, 1991). Eriophorum vaginatum, a tussock forming sedge, is the dominant plant, interspersed with a scattered mix of shrubs (Betula nana, Salix species, Vaccinium vitis-idaea), feather mosses (e.g. Hylocomium splendens and Dicranum elongatum), and sphagnum moss (Sphagnum rubellum). The mosses and shrubs mostly grow in the spaces between the 10 and 30 cm tussocks, spaces that are called ‘intertussock’. Tussock soil is always moist, and has an uneven organic layer 0–20 cm thick (Shaver and Chapin, 1991). All of the tussock samples were collected from the organic layer directly beneath E. vaginatum plants to a depth of approximately 15 cm. This soil is comprised primarily of decaying E. vaginatum roots. In intertussock soil, samples were collected underneath the mosses and shrubs between E. vaginatum tussocks. This organic soil consists of decomposing mosses, shrub fine roots and leaf litter. Another moist tundra form is shrub tundra, found in upland areas and along water tracks (Shaver and Chapin, 1991). The relatively tall (!1–2 m) shrubs Salix pulchra and B. nana dominate shrub tundra, with several other shrubs as lesser components (e.g. V. vitis-idaea). Shrub tundra is typically found on well-drained, gravelly soils, covered with a thin moss dominated organic soil 2–10 cm thick (Shaver and Chapin, 1991). Organic soil was sampled in the shrub community, to a depth of approximately 5–10 cm. Its appearance was similar to intertussock soil, but it contains less decaying mosses and more shrub litter and roots. The wet sedge tundra community occurs in flat, lowlying areas characterized by standing water throughout much of the summer, which results in high water contents and anaerobic soil conditions (Shaver and Chapin, 1991). Low stature (!20–30 cm) rhizomatous sedges such as Carex aquatilis and Eriophorum angustifolium predominate in these areas, with some Eriophorum scheuchzeri (Shaver and Chapin, 1991). The soil becomes wetter further north on the Arctic coastal plain, and wet sedge becomes the dominant plant community. Its soils are relatively thick peats (typically O30 cm) dominated by C. aquatilis fine roots. They typically thaw to a depth of 25–30 cm (Shaver and Chapin, 1991). The underlying permafrost prevents water drainage, keeping the soil flooded and cold. Soils in
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the wet sedge community were collected from the top 15 cm. 2.2. Sampling Soil samples from Weintraub and Schimel (2005) were analyzed. Sample collection began with spring thaw in June 2000, and was repeated every 1–2 weeks through August 2000, and on two dates in July 2001. We selected one general area to sample in each of the community types described. At each sampling we collected three cores from random locations within in each community; all cores were collected within 100 m of one another. Only organic soil was collected—mineral soil was excluded to avoid the influence of mineral type or parent material. Intertussock samples were collected as close to sampled tussocks as possible. Upon collection, samples were hand sorted (with gloves) to remove plants and debris. The soil samples were mixed by hand for several minutes to homogenize them. Samples from individual cores were kept separate as field replicates. 2.3. Protease activity The basic approach to the protease activity assay is to create a slurry in which the microbial uptake of amino acids is inhibited with toluene, so that protein breakdown by protease enzymes causes amino acids to accumulate in solution. The solutions are periodically sub-sampled, and the sub-samples are analyzed for total free amino acid (TFAA) concentration. Protease activity is calculated as the rate of amino acid accumulation over time in the slurry and expressed as mM amino acids gK1 dry soil hK1. We used the method of Watanabe and Hayano (1995) as modified by Lipson et al. (1999). Slurries were prepared by using 5 g wet soil in 40 ml 50 mM sodium citrate buffer in 125 ml specimen containers. The pH of the buffer was adjusted so that it was within 0.1 pH unit of the soil pH in each sample. Toluene (0.4 ml) was added to the slurries as a microbial uptake inhibitor. This procedure was named ‘actual’ protease activity, as the only substrate available to protease enzymes in these slurries was the native soil protein. Another version of the assay in which half of the buffer volume was replaced with a 0.6% casein solution was named potential protease activity. All slurries were kept at 5 8C, a typical tundra soil temperature (Weintraub and Schimel, 2005). Sub-samples (2.0 ml) were collected after 5 min, 4 and 6 h. Protease activity rates for the 0–4 and 4–6 h periods were calculated separately. A trichloroacetic acid/acetate buffer (2 ml) was added to each sub-sample to halt proteolysis (Lipson et al., 1999b). The sub-samples were then frozen until analysis. Means of the two rates from each sample are presented here. In some cases, there was a net immobilization of amino acids during one of the intervals, and the rate of protease activity for that interval was considered to be zero.
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When necessary, these declines in amino acid concentrations in the slurries were accounted for when calculating the rate for the following interval. Upon thawing for analysis, the sub-samples were centrifuged to remove any soil from the solution (IEC HN 30 min, w6000g). Then the supernatant was analyzed using a 96 well plate version of the ninhydrin colorimetric analysis for total amino acids (Lipson and Monson, 1998; Lipson et al., 1999b), with leucine as the standard. NHC 4 concentrations were not accounted for (Ladd and Butler, 1972; Watanabe and Hayano, 1995), as ammonification is mainly intracellular, and is inhibited by toluene (Lipson, pers. comm.). Also, even if extracellular amidases are active, the ammonium they produce will mostly be from glutamine and asparagine, and should be considered to be part of the amino acid N pool (Lipson, pers. comm.). Because there was a more than two order of magnitude difference in protein concentrations between the actual (w23 mg lK1) and potential assays (w2800 mg lK1), the difference between the two rates is indicative of the extent to which protease enzymes are limited by substrate availability. Little difference between the two, despite the large difference in protein concentrations, is an indication that native soil proteins were already saturating protease enzymes. 2.4. Soluble proteins Three 10 g (wet weight) sub-samples of each soil were shaken with 50 ml 0.1 M NaHCO3, an extractant mild enough to remove proteins from soil without lysing soil microbes (Ladd and Paul, 1973; Lipson et al., 1999b). The extractions were shaken on an orbital shaker table at w120 rev minK1 for 1 h in 100 ml specimen containers and were then vacuum filtered through Whatman GMF 2 mm filters. The extracts were frozen until analysis. To analyze these soil extracts for soluble proteins, we used Pierce Coomassie dry protein assay plates (Pierce Biotechnology Inc., Rockford, IL, Cat. no. 23296). The plates contain dried, stabilized Coomassie dye for the colorimetric determination of total protein concentration, using a Molecular Devices Vmax Kinetic microplate reader (Molecular Devices, Sunnyvale, CA) at 595 nm. Bovine serum albumin was used as a standard with a second order curve fit. 2.5. Statistics Systat version 10 (Systat Software Inc., Richmond, CA) was employed for statistical analyses, which were all conducted on untransformed data. Means of actual and potential protease activity were compared using T-tests. The means of both protease activity and soluble protein concentrations were compared by either soil type or date using one-way analysis of variance (ANOVA), followed by Tukey’s multiple comparisons test when the result from
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the ANOVA was significant. To compare these means by both soil type and date together two-way ANOVA was used. The strength of the relationship between two variables was examined by regression analysis. For the seasonal timecourse data, nZ9 time-points for protease activity in tussock, intertussock, and wet sedge soils, and nZ10 in shrub. For the soluble protein concentrations, nZ9 timepoints in tussock and intertussock, 7 in shrub, and 8 in wet sedge soils. Each time-point is the mean of values from typically three replicate samples collected on that date. The means of the replicates were used as the value for each timepoint so that the statistical analyses would not contain different numbers of values for different dates. Tests were considered significant if P!0.05 in all statistical tests unless otherwise noted. For regressions across the different soils the overall mean value for each soil type was used. When means for the different soils were used in the correlations, they had similar, but stronger patterns than those using individual sample values. This method reduces the influence of extreme values from individual sample replicates, and eliminates the problem of comparing different numbers of samples for different soil types in a regression. Regression slopes were significantly different than zero (P!0.05), unless otherwise noted.
3. Results Intertussock had, on average, the highest soluble protein concentrations, and actual and potential protease activity rates. Actual protease rates in intertussock were significantly higher than in tussock and wet sedge soils. Mean soluble protein concentrations in intertussock were significantly higher than in shrub soil, which had the lowest soluble protein concentrations. However, there was no significant difference between any of the soils in potential protease activity. In tussock and intertussock soils, actual and potential protease activities were equal (i.e. enzymes were substrate saturated) at all times except for one date in late July, and at the end of the growing season in intertussock, when proteolysis became substrate limited. Shrub soil protease activity had a different pattern, and was only substrate limited for a short period in late June into early July. Wet sedge protease activity was enzyme limited at the beginning and end of the growing season, but except for one date in early July, was substrate limited from late June to the beginning of August. In all soils, soluble protein concentrations paralleled potential protease activity, especially in June and July (Fig. 1). However, the seasonal dynamics of soluble proteins differed among the soils. In intertussock and tussock soils, there were peaks in late July, while the seasonal highs in shrub and wet sedge soils were in late
June, with another peak at the end of the growing season in wet sedge. Unlike potential protease activity, amino acid concentrations did not track soil protein, and the two often had opposing patterns, with soluble proteins frequently increasing after a decrease in amino acids (Fig. 1; see Weintraub and Schimel, 2005 for a detailed comparison of the soils’ amino acid concentrations).
4. Discussion The seasonal time-courses of actual and potential protease activities were similar to one another (Fig. 1; R2Z0.92), except for the short periods when potential rates peaked. This suggests that, contrary to our initial hypothesis, protein degradation was generally limited by enzyme activity, and only became substrate limited at the times when activity was highest (Fig. 1). Potential protease activity and soluble protein concentrations both peaked at the same times, when extractable N concentrations were declining (Fig. 1; data from Weintraub and Schimel, 2005). As a result of this pattern, both measures of protease activity were negatively correlated K with NHC 4 , NO3 , and TFAA, across all soils in our study 2 (R Z0.47–0.88). The question that arises is which way the causality goes in the relationship between increased soluble proteins and potential protease activity. On one hand, it is possible that increases in protein supply drove the increases in potential protease activity. On the other hand, it is also possible that soil microorganisms responded to low N availability by inducing protease production, and that the increases in soluble proteins resulted from faster breakdown of insoluble proteins into soluble fragments. In support of the protein supply argument, protein availability in soil is often described as being controlled by the turnover of microbes and fine roots (Nannipieri et al., 1979; Lipson et al., 1999a; Lipson and Nasholm, 2001; R.L. Sinsabaugh, pers. comm.) rather than by the solubilization of insoluble protein complexes. In the Alaskan arctic, soluble proteins peaked at the times of highest plant root growth (Shaver and Kummerow, 1992; Weintraub and Schimel, 2005). This raises the possibility that proteins in root exudates may have been responsible for some of the concentration increases (Saxena et al., 1999; Dakora and Phillips, 2002; Park et al., 2002). It is also possible that an increase in microbial turnover was mediated by root C inputs (Norton and Firestone, 1991, 1996; de Neergaard and Magid, 2001). Increased microbial turnover could either be a direct result of changes in substrate availability, or the result of an indirect effect, such as increased predation on soil microorganisms. Thus, root growth could stimulate protein availability even as amino acid and NHC 4 concentrations decreased due to higher plant N uptake.
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a Protease Activity
Actual Potential
2
2
1 0 3
1
(b)
b
Soluble Proteins
0 3
2
2
1
1
0
(c)
c TFAA
20 10
10
0
0 16 -Ju 01-Ju 16-Ju 31- 15-A 30Ju Au n ug l l l g
Shrub 3
0 20
n l ug ug ul ul -Ju 1-Ju 16-J 31-J 15-A 30-A 16 0
µg-N g–1 soil mg g–1 soil µM g–1 soil hr–1
3
Wet Sedge
(a) Actual Potential
Protease Activity
a 3
2
2
1
1
0 3
(b)
b
Soluble Proteins
0 3
2
2
1
1
0
µg-N g–1 soil mg g–1 soil µM g–1 soil hr–1
3
InterTussock
(a)
(c)
20
c
TFAA
0 20
10
10
0 n l ug ug ul ul -Ju 1-Ju 16-J 31-J 15-A 30-A 16 0
16 -J
30 16 01 31 15-A ug un -Jul -Jul -Jul Aug
0
µg-N g–1 soil mg g–1 soil µm g–1 soil hr–1
µg-N g–1 soil mg g–1 soil µM g–1 soil hr–1
Tussock
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Fig. 1. Maximum observed rates of potential (with added protein) and actual (no added protein) protease activity (panel a), soluble protein concentrations (panel b), and TFAA (panel c; Weintraub and Schimel, 2005) from the summer of 2000. The seasonal time-courses all show the sample replicate mean for each time-point (nZ3), and the error bars show the standard error of the mean.
Ladd and Paul (1973) and Lipson et al. (1999b) have shown that protease potential responds to elevated protein supply. Ladd and Paul (1973) observed a peak in protease activity following a peak and decline in soil microbial biomass, and concluded that the increase in protease activity was fueled by a flush of proteins released by the reduction in microbial biomass. Lipson and Na¨sholm (2001) also found the highest soluble protein concentrations and protease activity rates to occur immediately after a decline in soil microbial biomass, while Lipson et al. (1999) found that proteins released by microbial turnover at snowmelt stimulated protease activity. In support of the N-demand argument, on the other hand, microbial N acquisition is strongly regulated by N availability. For example, Smith et al. (1989) and Chrost (1991) both showed that proteolysis can be stimulated by N
deficiency, and depressed by adding N. Sinsabaugh and Moorhead (1994) found they could effectively model decomposition based on enzyme activities by assuming that microbes allocated resources based on nutrient demand, rather than on their supply. This suggests that as decomposers become N limited, allocation to N-acquiring enzymes increases. There are relatively recalcitrant peptide pools in soils such as glomalin (Zhu and Miller, 2003) and humus linked proteins (Yu et al., 2002), that can be degraded by protease, especially if there is substantial laccase/peroxidase activity, resulting in increased soluble protein concentrations (Sinsabaugh, pers. comm.). Plant growth is strongly N limited in tussock tundra (reviewed in Schimel et al., 1996), and recent research has found that microbial activity and growth are also N limited in tussock soils (Mack et al., 2004; Schimel, unpublished data). Thus,
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protease synthesis may be stimulated by high N demand in arctic tundra soils. Since protease can respond to either increases in available protein, or to reductions in N availability, this raises the question of how to determine which mechanism is operating in the Alaskan tundra. We propose that by examining the dynamics of soluble protein, potential and actual protease activity, and soil amino acids together, it may be possible to infer the mechanism. If increases in protease activity were driven by increases in protein supply, then enzyme activity would likely remain saturated by substrate availability, and actual rates would likely increase in step with potential. In this situation, one would also expect amino acid, and possibly inorganic N concentrations to increase in parallel with increases in protein supply and protease activity. For example, in alpine tundra soil, Lipson et al. (1999) found that a decline in microbial biomass at snowmelt caused a release of protein, which induced a pulse of protease activity, and that this was the only time that protease activity was not substrate limited. They also found that the peak in protein and protease was followed by an increase in amino acid and inorganic N concentrations. If, on the other hand, an increase in protease were driven by increased N demand and not by an increase in protein availability, the increase in the pool of enzymes would be large relative to substrate availability, causing greater substrate limitation to the enzymes, along with the decreased available N concentrations that induced N limitation and stimulated protease synthesis. Given the logic of this argument, then N limitation inducing protease synthesis is the most likely explanation for the increases in protease potential observed in these tundra soils. First, the times when soluble proteins peaked in our samples were the only times when protease activity was substrate limited, indicating that protease was increasing to a greater degree than protein supply. Second, potential protease activity peaked at a time when amino acid concentrations hit their lowest levels (Fig. 1). The fact that amino acid concentrations were low when protease activity (and therefore gross amino acid production) was high indicates that declines in TFAA (Fig. 1) were caused by increased plant and microbial uptake of amino acids, rather than a decline in their supply. Third, both actual and potential protease activity were poorly correlated with soluble protein concentrations (R2Z0.02–0.36, most not significant), but were strongly correlated with K2SO4-extractable dissolved organic nitrogen (DON), which is comprised largely of more recalcitrant protein complexes with humus or polyphenols (Yu et al., 2002; R2Z0.95 for actual protease activity; R2Z 0.76 for potential protease activity; DON data from Weintraub and Schimel, 2005). This is another indication that the increases in protease activity were not fueled by inputs of fresh proteins from microbial turnover or root exudation. These data suggest that the increases in soluble proteins (Fig. 1) were not the cause, but rather an effect of
Table 1 Reprinted from (Weintraub and Schimel, 2005), soil classifications (C.L. Ping pers. comm.), and mean N and C contents Soil type
Soil classification
%N
%C
Intertussock Shrub
Loamy, mixed, Typic Aquaturbel Loamy-skeletal, mixed, active, gelic Aquic Umbrorthel Loamy, mixed, Typic Aquaturbel Dysic, Typic Hemistel
0.5 1.3
42 38
1.6 2.0
33 37
Tussock Wet sedge
increased protease activity, resulting from the breakdown of insoluble proteins into smaller, soluble peptides (Table 1). Weintraub and Schimel (2005) concluded that the decreases in TFAA and other available N forms in these soils were driven by increased plant activity, because the declines coincided with the seasonal peaks in plant root growth and nutrient uptake. Taken together, these data suggest that the peaks in both microbial and plant amino acid demand coincide, and that both play a role in regulating N pool dynamics. The peak in proteolysis in July occurs at a time of high plant N uptake and root growth, suggesting that plant competition for N, and possibly root C inputs to the soil, result in microbial N limitation, and that this stimulates microbes to increase protease production. Acknowledgements This research was supported by the Andrew W. Mellon Foundation, and the ATLAS program. For their help with sampling and lab analyses, we thank Allen Doyle and Carl Mikan. For support in the field, we thank the Staff of Toolik Field Station. For the use of his equipment at Toolik Lake, we thank Dr Jeff Welker. References Asmar, F., Eiland, F., Nielsen, N.E., 1994. Effect of extracellular-enzyme activities on solubilization rate of soil organic nitrogen. Biology and Fertility of Soils 17, 32–38. Chapin, F.S., Bloom, A.J., Field, C.B., Waring, R.H., 1987. Plant responses to multiple environmental factors. Bioscience 37, 49–57. Chapin, F.S., Moilanen, L., Kielland, K., 1993. Preferential use of organic nitrogen for growth by a non-mycorrhizal arctic sedge. Nature 361, 150–153. Chrost, R.J., 1991. Environmental control of the synthesis and activity of aquatic microbial ectoenzymes. In: Chrost, R.J. (Ed.), Microbial Enzymes in Aquatic Environments. Springer, New York, pp. 29–59. Dakora, F.D., Phillips, D.A., 2002. Root exudates as mediators of mineral acquisition in low-nutrient environments. Plant and Soil 245, 35–47. de Neergaard, A., Magid, J., 2001. Influence of the rhizosphere on microbial biomass and recently formed organic matter. European Journal of Soil Science 52, 377–384. Finzi, A.C., Berthring, S.T., 2004. Organic nitrogen cycling in temperate forests of southern New England. Bulletin of the Ecological Society of America 89, 154. Giblin, A.E., Nadelhoffer, K.J., Shaver, G.R., Laundre, J.A., McKerrow, A.J., 1991. Biogeochemical diversity along a riverside toposequence in arctic Alaska. Ecological Monographs 61, 415–436.
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