There were two main objectives of this research. The first was to determine how slow vegetative succession influences soil properties. The second was to determine if soil characteristics influence early succession rates after forest clear-cut. To meet these objectives, a set of 24 research stands was selected on the H.J. Andrews and the adjacent Blue River watershed. Six sites exhibiting slow canopy closure 40 years post-harvest and 6 sites showing normal or expected rates of canopy closure. In addition, adjacent unharvested stands acted as pseudocontrols bring the total number of study stands to 24. Since elevation was thought to also influence recovery rates, half of the sites were at low elevations and half were at high elevations. Within each stand, three sample plots were established.
Site selection for this study was done in two phases; aerial photos, harvest records, TM imagery, DEM data and topographic maps were analyzed to identify candidate sites. From that cohort, the 12 sites used in this study were those which covered at least 5 hectares and where reasonable matches could be made between slow and expected sites at the same elevation. All sites were within the Blue River and Quartz Creek drainages of the Central Oregon Cascade Mountains in and adjacent to the H. J. Andrews Experimental Forest. Within each of these sites, three 250 m2 sample plots were established which were 17.42 m in diameter and separated by at least 100 m. All harvested plots were approximately the same age (40 years).
For the microbiological studies, five 4.8 x 10 cm cores were taken randomly within each of the three sample plots with all cores separated by at least 5 m. These cores were analyzed for the presence of ectomycorrhizal mats and then bulked into one large sample from which subsamples were taken for the laboratory studies.
Soil temperatures and litter depths were measured at each coring location using a calibrated dial thermometer to a depth of 10 cm and a ruler respectively.
The following measurements were made in the field: litter depth, soil depth, mineral soil respiration, ambient light, soil temperature and ectomycorrhizal mat characteristics. Field (mineral soil) 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. Light was measured with the Li-Cor photometer.
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).
The fraction dried weight and all laboratory measurements where taken on bulked soil samples that had been sieved through a 2mm mesh screen. 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 organic matter was measured by loss-on-ignition at 550 degrees C for 6 h after oven drying at 100 degrees C.
Duplicate denitrification potential measurements were made 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¡ÆC. The net N2O released over this 2-h period was used to estimate N2O production rates.
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 degrees 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). Duplicate aliquots and controls were run for all samples.
Griffiths, R.P., Homann, P.S., and Riley, R. (1998) Denitrification enzyme activity of Douglas-fir and red alder forest soils of the Pacific Northwest. Soil Biology and Biochemistry 30, 1147-1157.
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.
Zou, X., D. Binkley, and K. G. Doxtader. (1992) A new method for estimating gross phosphorus mineralization and immobilization rates in soils. Plant & Soil 147,243
Keeney, D.R., and Bremner, J.M. 1966. Comparison and evaluation of laboratory methods of obtaining an index of soil nitrogen availability. Agronomical Journal 58, 498-503.
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.
