• Sherri L. Johnson - Principal Investigator
  US Forest Service ;Pacific NW Research Station ;3200 SW Jefferson Way, Corvallis, OR, 97331, USA
  Phone: 541-758-7771
  Email: sherri.johnson2@usda.gov, sherri.johnson@oregonstate.edu
  URL: https://www.fs.fed.us/research/people/profile.php?alias=sherrijohnson
• Julia A. Jones - Principal Investigator
  Oregon State University;Department of Geosciences; Wilkinson Hall 104, Corvallis, OR, 97331-5506, USA
  Phone: (541) 737-1224
  Email: Julia.Jones@oregonstate.edu, geojulia@comcast.net
  URL: http://ceoas.oregonstate.edu/profile/jones/
  ORCID: http://orcid.org/0000-0001-9429-8925
• Matthew G Betts - Principal Investigator
  Department of Forest Ecosystems and Society; 201E Richardson Hall; College of Forestry; Oregon State University, Corvallis, OR, 97331
  Phone: (541) 737-3841
  Email: matt.betts@oregonstate.edu
  URL: http://www.fsl.orst.edu/flel/index.htm
• Michael P. Nelson - Principal Investigator
  Department of Forest Ecosystems and Society; 201K Richarson Hall; College of Forestry; Oregon State University, Corvallis, OR, 97331
  Phone: 541-737-9221
  Email: mpnelson@oregonstate.edu
  URL: http://www.michaelpnelson.com
  ORCID: http://orcid.org/0000-0001-6917-4752
• David Bell - Principal Investigator
  Email: david.bell@usda.gov, david.bell@oregonstate.edu
  URL: https://lemma.forestry.oregonstate.edu/about/david-bell

• The H.J. Andrews Experimental Forest is a living laboratory that provides unparalleled opportunities for the study of forest and stream ecosystems in the central Cascade Range of Oregon. Since 1980, as a part of the National Science Foundation Long Term Ecological Research (NSF-LTER) program, the Andrews Experimental Forest has become a leader in the analysis of forest and stream ecosystem dynamics.
• Long-term field experiments and measurement programs have focused on climate dynamics, streamflow, water quality, and vegetation succession. Currently researchers are working to develop concepts and tools needed to predict effects of natural disturbance, land use, and climate change on ecosystem structure, function, and species composition.
• The Andrews Experimental Forest is administered cooperatively by the USDA Forest Service Pacific Northwest Research Station, Oregon State University and the Willamette National Forest. Funding for the research program comes from the National Science Foundation (NSF), US Forest Service Pacific Northwest Research Station, Oregon State University, and other sources.

Data were provided by the HJ Andrews Experimental Forest research program, funded by the National Science Foundation's Long-Term Ecological Research Program (DEB 2025755), US Forest Service Pacific Northwest Research Station, and Oregon State University. National Science Foundation: DEB1440409

• Long-Term Ecological Research
  The Andrews Forest is situated in the western Cascade Range of Oregon, and covers the entire 15,800-acre (6400-ha) drainage basin of Lookout Creek. Elevation ranges from 1350 to 5340 feet (410 to 1630 m). Broadly representative of the rugged mountainous landscape of the Pacific Northwest, the Andrews Forest contains excellent examples of the region's conifer forests and associated wildlife and stream ecosystems. These forests are among the tallest and most productive in the world, with tree heights of often greater than 250 ft (75 m). Streams are steep, cold and clean, providing habitat for numerous aquatic organisms.

• Mineral soil samples were collected with a trowel to a depth of 10 cm and transported to the laboratory in an ice chest. Soils were stored at 15°C until the initiation of the analyses which was within 16 h of their receipt.
• Field forest floor respiration rates, soil temperature, and litter depth were all measured at the study sites. The 24 h forest floor respiration was measured using a standard soda-lime technique (Edwards, 1982) . At the end of the incubation period, a 10 mL headspace gas sample was removed from the incubation chambers before the soda-lime jars were removed. Headspace samples from sealed controls were used to subtract out ambient CO concentrations at the site prior to initiation of the incubation period. Headspace CO that was not absorbed by the soda lime during the incubation period was added to the total CO adsorbed into the soda lime. Headspace CO concentrations were measured on a Hewlett Packard model 5890 gas chromatograph fitted with Hewlett Packard model 3396 integrator. The GC was fitted with a flame ionization detector and a methanizer in series. The integrator was calibrated using known gas standards and the external calibration method.

• The other soil characteristics measured in this study were: soil organic matter, laboratory respiration, exchangeable ammonium, mineralizable nitrogen, water extractable organic carbon, denitrification potential, pH, bulk density, and moisture.
• In preparation for laboratory analysis, all soils except those used in the water extractable organic carbon measurements were sieved through a 2-mm sieve. Soil organic matter was measured by loss-on-ignition at 550°C for 6 h following oven drying at 100°C. Laboratory respiration rates were measured by adding 5 g field moist soil to a 25 mL Erlenmyer flask. After being sealed with a serum bottle cap, the flasks were incubated at 24°C for one hr. Zero-time headspace-CO concentration measurements were made after an initial 1 h preincubation period. Flasks were subsequently incubated for 2 h at 24 °C and the headspace analyzed again for CO. CO concentrations were assayed with a GC as described above. In the HO respiration experiments, the same procedure was followed as for laboratory respiration rates except 2.0 mL of sterile HO was added to the soils in the flasks; for SIR, 2 mL of sterile 1 mM glucose was added to the soils.
• Exchangeable NH concentration was determined by shaking 10 g field-moist soil with 50 mL 2 M KCl for 1 h (Keeney, 1982). After adding 0.3 mL 10 M NaOH to the slurry, NH concentration was measured with a Orion model 95-12 ammonium electrode (Orion Research Inc. Boston, Ma, USA). Mineralizable N was measured using the waterlogged technique (Keeney, 1982). Ten g of field-moist soil was added to 53 mL of distilled water in a 20 x 125 mm screw cap test tube and incubating at 40°C. After 7 days, 53 ml of 4 M KCl was added to the slurry and NH concentration was determined with the ammonium electrode. Mineralizable N was calculated from the difference between initial and final NH concentrations.
• Water extractable organic carbon (DOC) determinations were made using a Dormann Carbon Analyzer. DOC extractions were made on nonsieved samples. Five g of wet weight soil were slurried with 15 ml of water in 100 mL serum bottles and shaken for one hour at 24°C. Subsamples of the resulting slurry were centrifuged at 13,600 x g for 5 min. on a microfuge (Sorvall model MC12 V). Water-only blanks were run concomitantly to subtract out background organic carbon levels. Supernatants were extracted from the centrifuge tubes and frozen until analyzed.
• Just before analyzing the samples, they were thawed and vortexed to suspend the precipitate. A series of experiments conducted with identical subsamples had the following treatments: (1) not frozen, (2) frozen with vortexing, and (3) frozen without vortexing. It was found that treatments 1 and 2 produced the same result therefore we concluded that the freezing and subsequent resuspension did not bias the resulting measurements.
• Denitrification potential was measured using a method similar that used by Groffman and Tiedje (1989). Each reaction vessel (25 mL Erlenmeyer flask) contained 5 g of less than 2mm, field-moist soil. The flask was sealed with a rubber serum bottle stopper and purged with Ar to displace O in the headspace. After purging with argon, 2 mL of a 1mM solution of glucose and NO was added to the flask which were incubated at 25°C for one hr. This preincubation period was used because time series experiments on representative soils showed a lag in NO production during this period. The same experiments have shown linear NO production rates during the following 2-4 hr. After the preincubation period, 0.5 mL of headspace gas was removed from the reaction vessel and injected into a gas chromatograph fitted with an electron capture detector (Hewlett Packard model 5890 GC fitted with Hewlett Packard model 3396 integrator).
• Another headspace NO analysis was made after an additional two hr incubation at 25°C. The net NO released over this 2 h period was used to estimate NO production rates. Acetylene was not routinely added to the headspace to prevent the conversion of NO to N because randomly selected samples (10% of the total) were also assayed with a 10% acetylene atmosphere. There were no significant differences between NO production rates with and without acetylene.
• Soil pH was measured in 1:10 (soil: distilled water) slurries of oven-dried (100°C) soil. These slurries were shaken for 1 h prior to reading pH values with Sigma model E4753 electrode.

• Sampling frequency: once only

• Each of the three forest types; oldgrowth, thinned pole and pole stands (OG. T, and P) used in this study were organized into triads to reduce confounding elements while comparing treatments. Individual sites within each triad were matched by elevation, aspect and proximity. Three triads were analyzed in each of the three regions. At each site, soil samples were collected to a depth of 10 cm every 5 m along a 250 m transect.

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