• 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.

• 1. The sites were located approximately every 0.5 km along all accessible roads on the HJA.
• 2. During the first day, we installed in incubation chambers (3 gal plastic ice cream tubs fitted with a serum bottle septum for collection of headspace gas analyses) installed in the sodalime jars, took cores for bulk density and percent moisture and took soil and air temperature readings. All vegetation except moss was removed from the ground covered by the incubation chambers. Those chambers covering moss was noted as such.
• 3. On the second day, we took headspace gas samples and placed them in Vacutainers for subsequent analysis for CO concentrations using a gas chromatograph. The sodalime was retrieved, air and soil temperatures measured and cores were taken for subsequent chemical and biological analyses.

• Before sieving the soils, subsamples were taken for DOC and laboratory respirations. The soils were then sieved through a 2 mm sieve for subsequent analysis.
• Five g field-moist samples were placed in a 25 mL Erlenmeyer flask fitted with a serum bottle stopper. Three separate subsamples were run for each sample. To one, nothing was added, to another, 2 mL distilled sterile water was added and to a third 2 mL of 10-3 M sterile glucose (dextrose) solution.
• After the samples were preincubated for one hr at 19°C, the first headspace analyses was made. Two hours later after being incubated at the same temperature, another headspace analysis was conducted. The net increase in CO concentrations was calculated by subtracting the first reading from the second.
• The concentration of CO was measured on a gas chromatograph fitted with a methanizer and a flame ionization detector. The instrument was calibrated with a standard calibration gas at 0.997 %.
• Substrate induced respiration (SIR) was calculated by subtracting the CO released from the water amended soils from the CO released from the glucose amended soils.
• Five g of wet weight soil was diluted with 15 ml of water in a 100 mL serum bottle.
• The soils are then shaken for one hour at room temperature and then allowed to stand for one hour. One and one half ml of the slurry is removed and centrifuged in a microfuge for 5 minutes.
• After centrifuging, 0.5 ml of the supernatant was removed and placed in a 0.5 ml centrifuging tube. Two blanks containing just deionized water are also run with the set. The samples are then frozen.
• Before analysis the samples were thawed and vortexed to suspend the precipitate. The samples were analyzed for organic carbon using a carbon analyzer.
• Two 10 g field-moist soil samples are measured into Al weighing boats and dried at 100°C for 8 h before dried weight determinations were made. Percent moisture was calculated as ((wet wt.- dry wt)/dry wt)*100.
• One 1-2 g subsample of dried soil (100°C for 8-12 hr) was placed in a preweighed porcelain crucible and reweighed. The sample was fired to 550°C in a furnace for 6 h. After firing, the samples were weighed again. The resulting data were expressed as a percent of the weight loss relative to the initial dried wt of the sample.
• Five gdw soil was added to 50 ml deionized water. The soil slurries were shaken for a least one h before pH measurements were made.
• The pH meter was calibrated using pH 4.0 and 7.0 standards. The samples and standards were stirred while readings were made.
• Replicate 10 g field moist soils were added to 50 ml 2M KCl in 250 ml Erlenmeyer flasks and shaken for one h. in the presence of 0.4 ml 10M NaOH.
• The specific ion meter used in to make this measurements was calibrated using a set of ammonium standards made up in 2M KCl. An Orion ammonia electrode was use to make the measurements.
• Ten g of field-moist soil was added to a large screw-topped test tube and filled with deionized water, shaken with the cap on to make sure all soil was saturated. The tubes were then filled with water and sealed with no gas in the headspace.
• The tubes were then incubated at 40°C for 1 week.
• After the soils have incubated, they were placed into 250 ml flasks and an equal amount (50 ml) of 4M KCl was added. One third ml of 10 M NaCl were added to the flasks and they are then shaken on a rotary shaker for at least one h before the ammonium concentrations were measured.
• The ammonium concentrations were measured using the same technique used to measure extractable ammonium. The net mineralizable nitrogen was calculated by subtracting the extractable ammonium concentrations from the ammonium concentrations after 1 week incubation.
• The soils were prepared using the same procedure as that for laboratory respirations except nonseived soils were used and the headspace was purged with Ar for a minimum of 3 mins. at over 100 mL per min.
• The flasks were incubated for one hr at room temperature then one or two mL of a sterile solution containing 10-3 M glucose and nitrate was injected through the stopper.
• The soils were incubated for another hour at room temperature at which time the time 0 reading is taken.
• After two more hours, a second headspace analysis was made for nitrous oxide concentration using the electron capture detector.
• Eight oz wide mouth glass bottles with metal and plastic lids were filled with 30 g 6-12 mesh sodalime. These jars were baked at 100°C for 8 h, weighed and sealed before they were placed in the incubation chambers in the field.
• After they were opened over 24 h to collect the CO, they were sealed, returned to the lab and baked again with the lids off. The jar were then reweighed that weight gain recorded.
• After the incubation chambers had been in place for 24 h and the CO adsorbed, a 6", 20 gauge needle and a 60 mL syringe was used to remove a headspace sample. One syringe- full of gas was pulled into the syringe and then expelled back into the chamber and then a second syringe-full was extracted from the center of the chamber.
• All but 5 mL of the 60 were used to purge a 7 mL vacutainer. The final 5 mL were used to overpressure the tubes.
• The gas in the Vacutainer was analyzed for CO using the same GC set-up used to measure laboratory respiration rates.
• The total CO production was calculated by subtracting out the controls for both soda-lime and headspace CO (chambers that were sealed at the beginning of the incubation period). The total CO released was calculated using the volume of the chambers and the area trapped under the incubation chamber.
• Soil temperatures (to 5 cm) and air temperatures were take during both days in the field using a simple dial thermometer. The mean values are the average for the two days.
• Bulk density measurements were made by drying 4.7 x 10 cm cores at 100°C for 8 h.
• The site locations were estimated on a map of the HJA and digitized into the GIS system. From these location, elevations were estimated using data already in the system.

• Sampling frequency: once only

• During the same 24 hour period, all 182 sites studied were sampled. These sites represent the best possible geographical coverage from accessible roads at the HJA.

• Andrews Experimental Forest (HJA)
  W -122.26172200, E -122.10084700, N 44.28196400, S 44.19770400
  Altitude: 1631 to 1631 meter

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