Epilithic biomass and chlorophyll a (Entity 1): We collected seven replicate epilithic algae samples per reach at the beginning and at the end of the experiment. For each sample, we scrubbed known areas of four to eight rocks for a total of area of at least 50 cm2 and collected the slurry in a container. Samples were placed on ice in the field, frozen, and analyzed within one month of collection.
Vertebrates (Entity 2): We surveyed aquatic vertebrates of fishes and amphibians using 2-pass depletion backpack electrofishing at the beginning and end of the approximately 40 day field experiment in each reach. For each survey, we measured length (total length and additionally vent length for amphibians) and weight (g) of each individual aquatic vertebrate that was captured. We surveyed and removed aquatic vertebrates every 2-6 days in the depletion reach. All aquatic vertebrates removed from the depletion reach were added to the addition reach. We walked, but did not survey, the natural (control) and addition reaches when we did additional surveys in the depletion reach.
Nutrients (Entity 3): Two water samples were collected for nutrient characterization at the end of each reach at the beginning and at the end of the experiment into 250-mL, acid-washed, high-density polyethylene (HDPE) bottles. One of the bottles was filtered in the field through a GF/F filter. Both were frozen and taken to the lab.
Stream channel wetted widths (Entity 4): Widths were measured every five meters for at least the first 45 m of the reach using a measuring tape.
Stream habitat measurements (Entity 5): We characterized stream habitat by doing transects across the wetted width of the stream channel every five meters during week 1 of the experimental manipulation. We recorded water depth and substrata size every 20 cm across the stream-wetted width.
Canopy cover (Entity 6): Average percentage of canopy cover was estimated every five meters in the middle of the reach using a handheld concave densiometer (Lemmon 1956, Platts et al. 1987).
Fine benthic organic matter (Entity 7): We collected between five and seven replicate samples of fine benthic matter using a sampling cylinder of 15.24 cm diameter. The sediment was stirred to a depth of 6 cm and then, while swirling, a sample of the suspension for fine benthic organic matter (FBOM, < 1mm) was quickly collected in a bottle and placed in a cooler in the field then frozen, taken to the lab and analyzed within one month of collection.
Dissolved oxygen and specific conductivity (Entity 8): Dissolved oxygen, water temperature, and specific conductivity were recorded at the end of each reach using YSI 600 OMS-V2 multiparameter sondes equipped with a 6150 ROX optical dissolved sensor (Yellow Springs Instruments, Yellow Springs, Ohio).
Metabolism (Entity 9): We estimated stream metabolism following the open-channel method, by monitoring dissolved oxygen concentrations downstream of each reach for at least 5 days during week 1 and week 5 of the experiment. The gas exchange coefficient between the stream and the atmosphere was measured by performing a constant rate injection of propane gas and a conservative tracer immediately above the upstream section of the reach and collecting longitudinal samples when the co-injection reached plateau (for more details see Song et al. 2018, and Pennington et al. 2018). We collected 10 sample replicates at 5 sampling sites.
Streamflow (Entity 10): We calculated discharge, mean travel time, and mean velocity from the conservative-tracer breakthrough curves (Kilpatrick and Cobb 1985) with EC as a surrogate for concentration.
Macroinvertebrates (Entity 11): To identify food webs under manipulated aquatic vertebrate densities, we surveyed for macroinvertebrates using surber sampling at the end of the field experiment in five riffle habitats in the manipulated reaches: depletion and addition. Macroinvertebrates were preserved in 95% alcohol and taken to the lab for processing.
Song, C.; Dodds, W.K.; Ruegg, J.; Argerich, A.; Baker, C.L.; Bowden, W.B.; Douglas, M.M.; Farrell, K.J.; Flinn, M.B.; Garcia, E.A.; Helton, A.M.; Harms, T.K.; Jia, S.; Jones, J.B.; Koenig, L.E.; Kominoski, J.S.; McDowell, W.H.; McMaster, D.; Parker, S.P.; Rosemond, A.D.; Ruffing, C.M.; Sheehan, K.R.; Trentman, M.T.; Whiles, M.R.; Wollheim, W.M.; Ballantyne, F. 2018. Continental-scale decrease in net primary productivity in streams due to climate warming. Nature Geoscience. 11(6): 415-420. doi: 10.1038/s41561-018-0125-5
Pennington, Robert; Argerich, Alba; Haggerty, Roy. 2018. Measurement of gas-exchange rate in streams by the oxygen-carbon method. Freshwater Science. 37(2): 222-237. doi: 10.1086/698018
Lemmon PE. 1956. A spherical densiometer for estimating forest overstory density. For Sci. 2:314–320.
Platts WS, Armour C, Booth GD, Bryant M, Bufford JL, Culpin P, Jensen S, Lienkaemper GW, Minshall GW, Monsen SB, et al. 1987. Methods for evaluating riparian habitats with applications to management. Ogden (UT): USDA Forest Service, Intermountain Research Station. General Technical Report INT–221.
Epilithic biomass and chlorophyll a (Entity 1): After thawing, each sample was split into two aliquots; the aliquot for standing stock was sieved to remove large, non-algal particles, filtered onto pre-combusted and pre-weighed GF/F filters. These filters were then dried at 60 deg C, weighed, combusted at 500 deg C, and reweighed to provide estimates of both organic and inorganic fractions of standing stock. The second aliquot was analyzed for Chlorophyll a and phaeophytin concentration by filtering onto pre-combusted GF/F filters, and using the heated ethanol methodology (Sartory and Grobblear, 1984). All measurement of concentrations of pigments occurred using a Shimadzu UV-1700 UV-VIS spectrophotometer. Chlorophyll a pigment concentrations were corrected retroactively based on Parker et al. (2016).
Nutrients (Entity 3): The frozen water samples were sent to the Cooperative Chemical Analytical Laboratory (CCAL) at Oregon State University and analyzed for total phosphorus, total nitrogen, soluble reactive phosphorus, ammonium, nitrate, and dissolved organic carbon following standard methods (See http://www.ccal.oregonstate.edu/ for a list of analytical methods).
Fine benthic organic matter (Entity 7): After thawing, the samples were sieved to remove large particles and filtered onto pre-combusted and pre-weighed GF/F filters. These filters were then dried at 60 deg C, weighed, combusted at 500 deg C, and reweighed to provide estimates of the benthic organic matter.
Metabolism (Entity 9): Propane samples were analyzed using an Agilent 7890A Gas Chromatograph
Macroinvertebrates (Entity 11): Macroinvertebrates that were preserved in 95% alcohol in the field were taken to the lab to be cleaned, picked, and classified to the lowest taxonomic unit possible. In 2014, macroinvertebrates were classified by Shannon Claeson, PNW Research Station, and in 2015 and 2016, they were classified by Matt Whiles lab, Southern Illinois University.
Most of the analyses were run at the Collaboratory Lab at Oregon State University (https://water.oregonstate.edu/collaboratory)
Sartory, D.P. and Grobbelaar, J.U. (1984) Extraction of Chlorophyll a from Freshwater Phytoplankton for Spectrophotometric Analysis. Hydrobiologia, 114, 177-187. http://dx.doi.org/10.1007/BF00031869
Parker, S.P., Bowden, W.B. and Flinn, M.B. (2016), The effect of acid strength and postacidification reaction time on the determination of chlorophyll a in ethanol extracts of aquatic periphyton. Limnol. Oceanogr. Methods, 14: 839-852. doi:10.1002/lom3.10130
