It is likely that these meadows were originally established and maintained by aboriginal burning (Miller and Halpern, 1998). However, factors responsible for the current invasion are not known with certainty but climate change, fire suppression and termination of sheep grazing may all have played a role (Popenoe et al., 1992; Miller and Halpern, 1998). In a comprehensive study of mountain tree invasion in the Central Oregon Cascade Mountains, Miller and Halpern (1998) noted that in addition to the allegoric factors mentioned above, autogenic factors may also control this process (i.e. local influence of trees on the establishment of seedlings by altering microclimate). In addition to controlling moisture, the pioneer trees could also be altering soil properties resulting in increased seedling establishment.
Comparative studies between grasslands and forests have shown large differences in soil chemistry (Göceoðlu, 1988; Hart et al., 1992; Popenoe et al., 1992; Ross et al., 1996; Yakimenko, 1997); litter decomposition rates (Hunt et al., 1988; Köchy and Wilson, 1997) and food web compositions (Hunt et al., 1987; Ingham at al., 1989). Our study was designed to measure changes in both the chemical and biological characteristics of high-elevation mountain meadows as adjacent forests invade them. This includes an analysis of the transition zone to obtain a rough idea of which soil properties are most rapidly altered in response to the tree invasion.
At each of these positions a 4.7 x 10 cm soil core was taken for subsequent analysis. The samples were transported to the laboratory in an ice chest and subsequently stored at 15 degrees C until the initiation of analyses, usually within 16 h of their receipt. The following measurements were made in the field: litter depth, mineral soil respiration, soil temperature and ectomycorrhizal mat characteristics. Field (forest floor) respiration rates were measured with a nondispersive, infrared CO2 analyzer (Li-Cor, LI-6200). Measurements were made over a period of 1 min after the chamber gas reached ambient CO2 concentration. The instrument was calibrated on site against a known standard at each location. A Q10 adjustment was made for ambient soil temperature. Soil temperature was measured by electronic thermometers calibrated at 0 degrees C with ice water. The temperature probes were inserted into the mineral soil to a depth of 10 cm.
The distribution of ectomycorrhizal mats was determined visually in the field by inspecting the relative abundance of mats in 4.7 x 10 cm cores. Two distinct mat types were scored: (1) mats similar to those of the genus Hysterangium and (2) mats similar to those of the genus Gautieria. This approach has been used successfully in the past to document ectomycorrhizal mat distribution patterns in coniferous forests of the Pacific Northwest (Griffiths et al. 1996).
In preparation for laboratory analyses, all soils were sieved through a 2-mm sieve. Soil moisture was determined by drying duplicate 10 g field-moist sieved soils at 100 degrees C for at least 8 h. The percent soil moisture was calculated by dividing the difference between wet and dry samples and dividing that number by the dry wt., which was then multiplied by 100. Soil pH was measured in 1:10 (soil:distilled water) slurries of oven-dried (100 degrees C) soil. These slurries were shaken for 1 h prior to reading pH values with a Sigma model E4753 electrode. Soil organic matter was measured by loss-on-ignition at 550 degrees C for 6 h after oven drying at 100 degrees C.
Denitrification potential was measured using a method by Groffman and Tiedje (1989) as modified by us (Griffiths et al., 1998). Each reaction vessel (25-mL Erlenmeyer flask) contained 5 g of less than 2 mm, field-moist soil. Flasks were sealed with rubber serum bottle stoppers and purged with Ar to displace O2 in the headspace gas. After purging with Ar, 2 mL of a 1 mM solution of glucose and NO3- was added to each flask. Flasks were subsequently incubated at 25 degrees C for 1 h. This preincubation period was used because previous time-series experiments showed a lag in N2O production during this period. The same experiments have shown linear N2O production rates during the following 2-4 h (unpublished data). After the preincubation period, 0.5 mL of headspace gas was removed from the reaction vessel and injected into a gas chromatograph (GC) fitted with an electron capture detector (Hewlett Packard model 5890 GC, connected to a Hewlett Packard model 3396 integrator). The integrator was calibrated by the external calibration method with known gas standards. A second headspace N2O analysis was made after an additional 2-h incubation at 25 degrees C. The net N2O released over this 2-h period was used to estimate N2O production rates.
Extractable ammonium was determined by shaking 10 g of field-moist soil with 50 mL 2 M KCl for 1 h (Keeney and Nelson 1982), adding 0.3 mL 10 M NaOH to the slurry, and measuring ammonium concentration with an Orion model 95-12 ammonium electrode (Orion Research Inc., Boston, MA). Mineralizable N was measured by the water-logged technique of Keeney and Bremner (1966). For each analysis, 10 g of field-moist soil were added to 53 mL of distilled water in a 20 x 125 mm screw-cap test tube, and incubated at 40 degrees C for 7 d. Then 53 mL of 4 M KCl were added to the slurry, and ammonium concentration was determined with the ammonium electrode. Mineralizable N was calculated as the difference between initial and final ammonium concentrations.
Beta-glucosidase activity was determined by the spectrophotometric assay of Tabatabai and Bremmer (1969), as modified by Zou et al. (1992). One mL of 10 mM p-nitrophenyl b-D glucopyranoside substrate was added to duplicate 1-mL subsamples containing a soil slurry (1 gdw in 1 mL deionized H2O). The tubes were shaken and then placed with duplicate controls without substrate in a 30°C water bath for 2 h. After incubating, 1 mL of 10 mM p-nitrophenyl b-D glucopyranoside was added to the controls, and all reactions were immediately stopped by the addition of 2 mL of 0.1 M tris[hydroxymethyl]aminomethane at pH 12.0. The mixtures were centrifuged for 5 min at 500 x g. From the supernatant, 0.2 mL was diluted with 2.0 mL deionized water. The optical density was measured at 410 nm, and a standard curve was prepared from 0.02 to 1.0 micro-mol/mL p-nitrophenol (pNP).
Groffman, P.M., and Tiedje, J.M. 1989. Denitrification in north temperate forest soils: relationships between denitrification and environmental factors at the landscape scale. Soil Biology & Biochemistry 21, 621-626.
Tabatabai, M. A., and J. A. Bremner. (1969) Use of p-nitrophenyl phosphate for assay of soil phosphatase activity. Soil Biology & Biochemistry 1,301 - 307.
Keeney, D.R., and Nelson, D.W. 1982. Nitrogen-inorganic forms. In Methods of soil analysis. Edited by A.L. Page, R.H. Miller, and D.R. Keeney. American Society of Agronomy, Madison, Wis. pp. 643-698.
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