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Methods for Experiment 247 -

Nutrient Network Experiment design and protocols

The NutNet study is a completely randomized block (environmental gradient) design with three blocks and 10 plots per block (N = 30 total units/site). Each experimental unit is a 5 by 5 m plot that is separated by at least 1-m walkways. Each plot is divided into four equal-sized subplots: one dedicated to core sampling, one to additional site-specific or subnetwork studies and the last two for the future network-level research. Core Sampling. In each plot, the core sampling 2.5 by 2.5 m subplot is divided into four 1 by 1 m permanent subplots, surrounded by a 0.25-m buffer. Within the core-sampling subplot, one 1-m2 subplot is permanently marked for annual plant composition sampling; the other three are used for destructive biomass sampling. Core annual sampling includes clipping of total above-ground biomass of all plants rooted within two 0.1-m2 strips (10 by 100 cm) for a total of 0.2 m2. These are sorted to live and dead (or further, e.g. forb, grass, moss at many sites), dried at 60 degrees C to constant mass and weighed to the nearest 0.01 g. Leaves and current years woody growth are collected from shrubs and subshrubs. Light availability above and at ground level below the canopy is measured in the core subplot using a linear 1-m bar (e.g. Apogee Instruments, Inc., Logan, UT, USA). Areal cover is estimated to the nearest 1% for each species rooted in the core subplot; cover estimates include woody overstorey, litter, bare soil, rock and animal activity (e.g. digging). All core measurements are collected from all plots, annually at peak biomass. Two 2.5 cm diameter by 10 cm depth soil cores, free of litter and vegetation, are collected from each plot prior to initiation of the experiment (Y0) and 3 years after treatment initiation (Y3). Soils from each plot are composited, homogenized, air-dried and shipped to a single laboratory for analysis and long-term storage. Samples are assayed for % total C and % total N, extractable soil phosphorus, potassium and micronutrients, soil pH, soil organic matter and soil texture. The experimental treatments are applied at the scale of the 5 by 5 m plots, as follows: Fertilization treatments. Three nutrient treatments (N, P and K plus micronutrients), each with two levels (control, added), are crossed in a factorial design, for a total of eight treatment combinations per block, to test multiple nutrient limitation on plant composition and ecosystem function. Nutrient addition rates and sources are: 10 g N m-2 year-1 as timed-release urea [(NH2)2CO], 10 g P m-2 year-1 as triple-super phosphate [Ca(H2PO4)2], 10 g K m-2 year-1 as potassium sulphate [K2SO4] and 100 g m-2 of a micronutrient mix of Fe (15%), S (14%), Mg (1.5%), Mn (2.5%), Cu (1%), Zn (1%), B (0.2%) and Mo (0.05%). N, P and K are applied annually; micronutrients are applied once at the start of the experiment to avoid toxicity. In addition to the standard NutNet protocol, e247 includes an additional low Nitrogen gradient (1 gram Nitrogen per meter squared per year and 5 grams Nitrogen per meter squared per year in addition to the standard 10 grams Nitrogen per meter squared per year). Fencing treatments. A fencing treatment is crossed with the control and NPK treatments to assess the interactive effects of fertilization and food web manipulation on plant composition and ecosystem function. The 230-cm-tall fences restrict access by mid-to-large-sized above-ground mammalian herbivores (>50 g). The lower 90 cm is surrounded by 1-cm woven wire mesh (hardware cloth) with a 30-cm outward-facing flange stapled to the ground to exclude digging animals (e.g. rabbits, voles), although not fully subterranean ones (e.g. gophers, moles). The upper fence is composed of four strands of tensioned wire strung at equal vertical intervals. The experimental design and sampling are replicated at grassland sites around the world. Most sites are contributing pre-treatment and all experimental data (full experiment); however, some sites have contributed only pre-treatment data (observational), and a few are applying only the nutrient addition treatments (nutrients only). Plots at Cedar Creek were established in 2007. See the following for further information: Borer, E.T. et al.; Finding generality in ecology: a model for globally distributed experiments; Methods in Ecology and Evolution; 2014; 5: 65 - 73.; doi: 10.1111/2041-210X.12125 2014

acue247 - Aboveground Standing Crop Biomass

Aboveground Standing Crop Biomass

Aboveground standing crop will be estimated destructively by clipping at ground level all aboveground biomass of individual plants rooted within a 0.2 m2 (two 10 x 100 cm) strips. Biomass will be clipped within the the 1-m2 subplots designated for destructive sampling within the core sampling subplot. Location of the quadrats should be noted or marked permanently to prevent resampling during the duration of the study. For shrubs and subshrubs rooted within the quadrat, leaves and current year?s woody growth should be collected. Standing crop should be separated into the following categories: previous year?s dead, current year?s bryophytes, and current year?s vascular plant. If time permits, it would be highly valuable to separate biomass into the following six categories: 1. previous year?s dead, 2. current year?s bryophytes, 3. current year?s graminoid (grasses, sedges, rushes), 4. current year?s legumes, 5. current year?s non-leguminous forbs, 6. current year?s woody growth. All biomass should be dried at 60?C for 48hrs prior to weighing to the nearest 0.01 g.

acze247 - Plant Species Composition percent cover

Plant Species Composition percent cover

Prior to initiation of the experiment, percent aerial cover will be estimated in one permanently marked 1-m2 subplot, one within the core-sampling subplot. Aerial cover will be estimated for each plant species separately using a modified Daubenmire method (Daubenmire 1959), in which cover is estimated to the nearest 1% percent for each species rooted within the plot (cardboard cutouts can be used to facilitate estimation). Percent cover also should be estimated for woody overstory, litter, bare soil, animal diggings/disturbance, and rocks if present. Note that total cover will typically exceed 100% because species cover is estimated independently for each species. Within-season sampling frequency will need to be adjusted for individual ecosystems based on the phenology of the component species in order to capture the maximum cover of each species, which will be used in subsequent analyses. For example, in the tallgrass prairie, species composition will be measured in the spring (late-May) and again in the fall (late-Aug) to capture maximum relative cover of early-season C3 forb and grass species and late-season C4 forb and grass species, respectively.

adae247 - Light Availability

Light Availability

Light availability will be measured using a light meter (e.g., 1-m length Decagon Ceptometer if possible) capable of integrated measures of photosynthetically active radiation (PAR, mmol m-2 sec-1). Light availability will be measured at the same time and in the same 1-m2 subplot used for the species composition measurements. Light readings must be taken on a cloudless day as close to solar noon as possible (i.e., 11 am to 2 pm). For each subplot, two light measurements at ground level (at opposite corners of the 1-m2 plot, diagonal to each other) and one above the canopy will be taken. Light availability will be calculated as the ratio of PAR below and above the canopy. Note: Light was not measured at the Cedar Creek site in 2011

adbe247 - 2007 pre-treatment soils pH, nutrients, texture

2007 pre-treatment soils: pH, nutrients, texture

Prior to initiation of the experiment, soil cores will be collected during the growing season from all of the plots. For each plot, collect two to three soil cores (soil corer - 2.5 x 10 cm) from each of the 2.5 x 2.5 m subplots (in areas designated for destructive biomass sampling). Litter and vegetation should be removed from the soil surface before collecting each sample. Composite and homogenize these sub-samples into a single sample for each 5x5 m plot (total of 30 roughly 500 g samples). All soil samples should be double bagged in paper and allowed to air dry. Label each bag (with permanent marker, Sharpie preferred) with the following information: date of collection, name of collector, name of sampling site, and block/plot/treatment identification. PERCENT CARBON and NITROGEN, by mass, as determined by COSTECH Analytical Elemental Combustion System 4010 (ESC 4010)[Valencia, CA USA] P, K, Ca, Mg, S, Na, Zn, Mn, Fe, Cu, B as determined by Melich-3 analysis (A&L Labs, Memphis, TN USA) pH as determined by water pH meter with soil in 1:1 soil:water suspension (A and L Labs, Memphis, TN USA)as determined by mixing known PERCENT SAND, SILT, CLAY as determined by mixing known amount of sample with water; allowing different particle types (sand, silt and clay) to settle out over time; measuring each layer to calculate what portion it is of the whole sample. (A and L Labs, Memphis, TN USA) PRE-TREATMENT SOIL CLASSIFICATION determined by relative fractions of sand, silt, and clay (A and L Labs, Memphis TN USA)

aeme247 - Multi-site grassland plant biomass, species richness and light (PAR)


This dataset contains multi-year data (up to 5 years of data at each site) from 40 global Nutrient Network sites, including five USA LTER sites, published in: "Borer et. al.; Herbivores and nutrients control grassland plant diversity via light limitation; Nature.2014; 508(7497):517-520., doi:10.1038/nature13144" Corresponding author: Elizabeth T. Borer Email:

Compiled data methods

We used an experiment replicated at 40 sites on 6 continents to test the hypothesis that herbivores mediate species losses caused by nutrient addition by increasing ground-level light, particularly in eutrophic and highly productive systems. We manipulated herbivores and nutrients using a factorial experiment (nutrient addition x exclusion of herbivores > about 50 g, see Methods Borer et. al. 2014a and 2014b for details) replicated in 40 sites dominated by herbaceous plants, spanning broad environmental gradients of productivity (114 to 1,976 g per meter squared per year), precipitation (mean annual precipitation from 224 to 1,898 mm per year), temperature (mean annual temperature from 0 to 22.1 degrees C), and soil nitrogen (mean percentage of soil N from 0.018 to 1.182 percent). In each plot, we measured local-scale responses of productivity, light and the number of plant species (diversity) using standard methods (Borer et. al. 2014b). We also examined site-level covariates including precipitation, temperature, herbivory intensity, soil nitrogen and atmospheric-nitrogen deposition rates. Although most sites provided 3 years of data, a subset of sites contributed 4 years of post-treatment data, and a few sites, established later, provided only 1 or 2 years of data (Extended Data Table 1 Borer et. al. 2014a). Method references cited: Borer et. al.; Herbivores and nutrients control grassland plant diversity via light limitation; Nature, doi:10.1038/nature13144 2014a Borer et. al.; Finding generality in ecology: a model for globally distributed experiments; Methods in Ecology and Evolution; 2014b; 5: 65?73.; doi: 10.1111/2041-210X.12125


Soil Nitrogen analyses: COSTECH Analytical Elemental Combustion System 4010 (ESC 4010)[Valencia, CA USA]

aeve247 - Soil organic matter responses to nutrient enrichment in the Nutrient Network

Sampling and Lab methods

Soil sampling Soils were sampled from nutrient addition plots at five participatory sites of the Nutrient Network. The Nutrient Network is a collaborative, global network of experiments established to investigate the effects of multiple nutrient additions, including N, on ecosystem processes in grasslands. Participatory sites are located across the globe and follow standard protocols for sampling and analysis (Borer et al. 2014). Soil samples were collected in July and August of 2012. Three cores (5 cm diameter and 10 cm deep) were sampled from each plot and composited across the full factorial of nutrient treatments; a fourth core was sampled from the control and N addition treatments for root analyses. Samples were kept on ice or in the refrigerator for a maximum of 6 days until processed in the lab. A subsample of composite soil from each plot was sieved to 2 mm for chemical and biological analysis and 8 mm for soil aggregate isolation. Fresh, 2 mm-sieved soil was used to mea-sure gravimetric soil moisture, microbial respiration, microbial biomass and net N mineralization. Air-dried, 2 mm-sieved soil was used to measure total soil % C and % N by combustion (Costech ESC 4010 Elemental Analyzer, Valencia, California, USA), soil pH (1:1 soil:water slurry method), and particulate organic matter (POM) C and N via density flotation (method detailed below). Additionally, soil texture was measured on air-dried, 2 mm-sieved soil from the control plots using the hydrometer method and sodium hexametaphosphate as the dispersing agent (Ashworth et al. 2001). Soil sieved to 8 mm from the control and N addition plots was air-dried and used to measure water-stable soil aggregates. Analyses: decomposition of unoccluded SOM A subsample of fresh, 2 mm sieved soil from each plot was placed in a 120 ml specimen cup and soil moisture was adjusted to 70 percent field capacity. field capacity was calculated separately for each site by pulling 20 kPa pressure on saturated soil. Microbial respiration rate (mg C g soil-1 day-1) was determined at least 17 times during the 380-day laboratory incubation. For each respiration rate measurement, the specimen cups were placed inside 1 L Mason jars and sealed for either 24- or 48-hour intervals. The CO2 concentration in the airtight jars was measured at the beginning and end of each interval using an infrared gas analyzer (LICOR 7000). When not being measured, specimen cups were covered with gas-permeable, low-density polyethylene film. Throughout the incubation, soil samples were maintained at 70 percent field capacity and kept at 20 degrees C in the dark. We calculated cumulative C respired (mgCgsoil-1) during the incubation by averaging the respiration rate between adjacent measurement dates and multiplying by the interval between them, then summing the amount of C respired in between each rate measurement. We assessed the effects of N addition on microbial biomass C and N at the start of the respiration incubation using chloroform fumigation extraction (Brookes et al. 1985). Briefly, replicate fresh, 2 mm-sieved soil samples were extracted with 0.5 M K2SO4 prior to and following chloroform fumigation under vacuum for 5 days. Following filtration, extracts were analyzed for total organic C and total N (Shimadzu TOC-V, Shimadzu Corpora-tion, Kyoto, Japan). Soil microbial biomass C (MC) and N (MN) were calculated as: MC = EC/kEC and MN = EN/kEN, where EC is the difference between extractable C in the fumigated and unfumigated samples, EN is the difference between extractable N in the fumigated and unfumigated samples, kEC is the C extraction efficiency coefficient, and kEN is the N extraction efficiency coefficient. We used the standard extraction efficiency coefficients of 0.45 (kEC) and 0.54 (kEN) from the literature (Brookes et al. 1985; Beck et al. 1997). Analyses: aggregate-occluded and mineral-associated SOM Briefly, air-dried, 8 mm-sieved soil subsamples from the control and N addition treatments only were wet sieved with a 2 mm sieve for 2 min each to isolate large macro-aggregates ([2000 lm). Soil that passed through the sieve was wet-sieved with a 250 lm sieve to isolate small macro-aggregates (2000 - 250 lm). finally, the remaining material was wet-sieved with a 53 lm sieve to isolate micro-aggregates (250 - 53 lm) and mineral-associated SOM (\53 lm). During wet sieving, floating organic matter was removed so we could test for N effects on C that was occluded within each aggregate fraction. The isolated fractions were dried at 105 degrees C for12 h, followed by 60 degrees C for 48 h. Fractions were weighed and analyzed for C and N concentration (Costech ESC 4010 Elemental Analyzer, Valencia, California, USA) and used to determine percentage of whole soil total C and N contributed by each fraction. The large macro-aggregate, small macro-aggregate, and micro-aggregate fractions were used to evaluate H2 (aggregate-occluded SOM), while the smallest size fraction informed H3(mineral-associated SOM). Directly following collection, the additional intact core sampled from the control and N treatment plots was washed in wire mesh tubes (0.28 mm mesh) in a rotating elutriator (Wiles et al. 1996)until allsoil was removed (*3 h). Remaining material was suspended in water and roots were captured with fine sieves and hand-picking. Root crowns were not considered root biomass and removed. Once free of soil, roots were dried at 65 degrees C overnight and weighed to calculate dry root biomass per unit area. Colonization of root tissue by arbuscular mycorrhizal fungi was determined by the point intercept method. Roots were removed from soil cores by washing gently with water over a 53 lm sieve. Cleaned roots were stained with Trypan Blue and stored in a 1:1:1 (vol) solution of glycerin:lactic acid:water at 4 degrees C. Roots were spread in a petri dish marked with 13 mm square grid and examined at 940 magnification to determine presence of fungal structures (hyphae and/or vesicles) at each root-grid line intersection. One hundred intersects were counted for every sample to determine the proportion of root tissue colonized, and each sample was counted twice to ensure reproducible results. Seven root samples were not prepared for mycorrhizal analysis and, consequently, are not included in the statistical analyses. Results from these data and full statistical analysis used for decay and respiration are presented in: Riggs, Charlotte E.; Hobbie, Sarah E.; Bach, Elizabeth M.; Hofmockel, Kirsten S.; Kazanski, Clare E.; (2015) Nitrogen addition changes grassland soil organic matter decomposition. Biogeochemistry, DOI: 10.1007/s10533-015-0123-2 Methods cited: Ashworth J, Keyes D, Kirk R, Lessard R (2001) Standard procedure in the hydrometer method for particle size analysis. Commun Soil Sci Plant Anal 32:633 to 642. doi:10.1081/CSS-100103897 Borer ET, HarpoleWS, Adler PB et al (2014) finding generality in ecology: a model for globally distributed experiments. Methods Ecol Evol 5:65 to 73. doi:10.1111/2041-210X.12125 Brookes PC, Landman A, Pruden G, Jenkinson DS (1985) Chloroform fumigation and the release of soil nitrogen: a rapid direct extraction method to measure microbial biomass nitrogen in soil. Soil Biol Biochem 17:837 to 842. doi:10.1016/0038-0717(85)90144-0 Beck T, Joergensen RG, Kandeler E et al (1997) An inter-laboratory comparison of ten different ways of measuring soil microbial biomass C. Soil Biol Biochem 29:1023 to 1032. doi:10.1016/S0038-0717(97)00030-8

afje247 - Soil nutrient analysis

Soil nutrient analysis

In 2012 four soil cores, 2.5 cm in diameter and 10 cm deep were taken from the 10 by 200 cm biomass clip strip after biomass and letter were removed. Samples were sieved and homogenized prior to analysis.

afre247 - Mechanisms driving the soil organic matter decomposition response to nutrient enrichment

Sampling and Lab Methods

Soil sampling and processing In August 2014, we collected soil samples from three multi- factorial nutrient addition experiments in the U.S. Central Great Plains: Cedar Point Biological Station (Ogallala, Nebraska; 41.20, -101.630); Konza Prairie Biological Station (Manhattan, Kansas; 39.070, -95.580); and Shortgrass Steppe (Nunn, Colorado; 40.820, -104.770). At each plot, six 0e10 cm cores (2 cm diameter) were sampled and kept on ice or in the refrigerator until processed in the lab. Within four days, soils samples from each plot were composited and sieved to 2 mm. Fresh, sieved soil was subsampled for further analysis (see below). Air-dried, sieved soil was used to measure total soil % C and % N by combustion (COSTECH ESC 4010 Elemental Analyzer, Valencia, CA, USA) and soil pH (1:1 soil:water slurry). Total soil C is equivalent to total soil organic C. Fine root samples from a subsample of fresh soil were collected via flotation and straining with a 250 mm sieve, along with roots captured on the 2 mm sieve; roots were cleaned of soil debris, dried, and analyzed for % C and % N by combustion (COSTECH ESC 4010 Elemental Analyzer, Valencia, CA, USA). Microbial biomass C and N and dissolved organic C Microbial biomass C and N were analyzed using a chloroform fumigation extraction procedure. Briefly, fresh, 2 mm sieved soil was extracted with 0.05 M K2SO4 and filltered. A replicate soil sample was fumigated with chloroform in a vacuum for 72 h, extracted with 0.05 M K2SO4 and filltered (Whatman No. 42; 2.5 mm pore size). Filtered extracts were analyzed for total dissolved organic C and total dissolved N (Shimadzu TOC-V, Shimadzu Corporation, Kyoto, Japan). Microbial respiration and decomposition parameters We measured microbial respiration and decomposition rates during a long-term laboratory incubation. Before we initiated the laboratory incubation, a 50 g subsample of fresh, sieved soil from each plot was adjusted to 70% field capacity and pre-incubated for 6 h at 20 0C in the dark. Field capacity was calculated separately for each site by pulling 20 KPa pressure on saturated soil. Microbial respiration rate (mg CO2eC g-1 soil day-1) was calculated by measuring the accumulation of CO2 in airtight 1 L Mason jars during 24e48 h intervals on days 1, 3, 6, 9, 12, 18, 24, 31, 38, 45, 60, 74, 96, 124, 152, 180, 208, and 236 of the incubation. We measured CO2 concentration inside the jars at the start and end of each interval by analyzing air samples collected via syringe with an infrared gas analyzer (LICOR 7000). When not being measured, soil samples were covered with gas-permeable, low-density poly- ethylene film and kept at 20 0C in the dark. Soil samples were maintained at 70% field capacity throughout the incubation. Microbial carbon use efficiency: isotope addition experiment Six replicate subsamples from each plot were prepared (10 g fresh, sieved soil samples) and brought to 70% field capacity with the addition of a C substrate solution (60 ??g C g???1 dry soil). Soils were placed in airtight 1 L Mason jars and respired CO2 was sampled at 0 and 24 h using an infrared gas analyzer (LICOR 7000). At both sampling times, 16 ml gas samples were collected by syringe and stored in 12 ml evacuated exetainers (Labco Limited, Lampeter, Wales, United Kingdom) for 17 weeks prior to analysis. Carbon dioxide, and its atom % 13C, were measured by gas chromatograph-isotope ratio-mass spectrometer (University of California, Davis, Stable Isotope Lab). Results from this data and full methods are published in: Riggs, C. E. and Hobbie, S. E. (2016). Mechanisms driving the soil organic matter decomposition response to nitrogen enrichment in grassland soils. Soil Biology and Biochemistry 99: 54-65. 2016

ahne247 - Plant and soil organic matter responses to ten years of nutrient enrichment in the Nutrient Network


Sampling Soil and plant samples were collected from nine temperate U.S. grassland sites that are part of the Nutrient Network. Following the Nutrient Network site name codes, sites included BNCH, CDPT, CDR, KZA, LOOK, SEV, SGS, TMPL, and TREL. Three blocks were included for each site except for CDPT, CDR, and SEV where 4 blocks were included. Aboveground plant tissue chemistry was analyzed on plant material collected in 2017 for all sites except for SGS and TMPL, which were sampled in 2016 ANPP is for 2017 for all sites except for TMPL, where ANPP was not available for 2017 so 2016 data were used instead. At KZA, soils were shallow and 15-30cm soil samples were not taken. In plots where soil depth less than 15cm, approximate soil depth was measured with a standard measuring tape and recorded. For soils from sites suspected to have high level of carbonates in the soil, soil samples were first fumigated with 12M HCl prior to percent Carbon analysis via combustion. All data are from Nutrient Network plots where `anthropogenic` category equals not restored/created and `exclose` equals control (not fenced).

Biomass protocols

ANPP was measured by site leaders at each site and reported to Nutrient Network database. 10 cm x 100 cm strip clipped above soil during peak biomass. Biomass is dried and weighed. BNPP (root ingrowth) measured using root ingrowth technique. Three ingrowth cores (0-15 cm depth, 5 cm diameter) were installed in fall 2016 (2:1 sand:soil) and removed in fall 2017 by slicing around the outside of the plastic mesh and pulling them from the soil. The soil from each ingrowth core was sieved separately to 2 mm and the roots removed. The roots were kept in separate bags, dried, and weighed. The dry root weights from the ingrowth cores were then averaged in each plot to estimate BNPP. Plant biomass fiber content determined using series of extractions in the order of: NDF (Neutral Detergent Fiber = soap + water), ADF (Acid Detergent Fiber 1 N sulfuric acid + soapy water), then ADL (Acid Determined Lignin = 72 percent sufuric acid). Root standing stock (RSS) measured at the time of root ingrowth installation. Three soil cores were extracted in each plot (0-15 cm depth) using a 5 cm diameter soil core. Soil cores were sieved separately using 2 mm sieve, and roots were removed. Root were dried, weighed, and dry root weights were averaged in each plot to estimate RSS.


Soil percent Carbon and percent Nitrogen were analyzed by combustion using COSTECH ESC 4010 Elemental Analyzer, Valencia, CA, USA. Base cation concentrations were measured by inductively coupled argon plasma (ICP) optical emission spectrometry (OES) using a Thermo-Scientific iCAP 7400 IC spectrometer. Oxalate-extractable Al and Fe were also measured by ICP.

Soils protocol

0-15 cm soils were sampled with 5cm diameter soil core; 15-30 cm soils were sampled with a 2.54 cm diameter core. Bulk density was measured by weighing air-dried soil from the soil core both prior to sieving (preDb) and following passing the soil through a 2mm sieve (postDb). Soil pH was measured in a 1:2 soil to water slurry. Soils were fractioned using a composite density and size fractionation procedure. Sieved and air-dried soils were shaken with DI water for 15 minutes, then centrifuged for 15 minutes. After decanting the supernatant (dissolved organic matter), each soil sample was mixed with sodium polytungstate (1.85 g cm-3), shaken lightly for 18 hours and then centrifuged at 3400 rpm for 30 minutes. The suspended light POM was aspired and washed, collected on a 20 micrometer nylon mesh filter, and dried at 60degreesC. The remaining sample was then washed with deionized water iteratively, vortexed to suspend, centrifuged for 20 minutes, and decanted three times, or until the supernatant was clear. Finally, each sample was passed through a 53 micrometer sieve to separate into heavy POM (greater than 53 micrometer) and MAOM (less that 53 micrometer). Each fraction was dried at 60 degrees Celsius, ground to a fine powder, and analyzed for percent Carbon and percent Nitrogen via combustion. Base cation concentrations were measured on sieved (2mm), air-dried soil. Soils were extracted with 10 mM NH4HCO3 (1:5 w:v of soil:acid), shaken for 2 hours and centrifuged at 4000g for 10 minutes. The supernatant of each sample was decanted onto 0.2 micrometer filters pre-rinsed with extractant. The extractant was subsequently acidified with 5percentnitric acid for storage at 4 degrees Celsius. Oxalate extractable Al and Fe was measured by first shaking 0.5 g air-dried soil with 30 mL of an oxalate solution (4:3 ratio by volume of 0.2 M ammonium oxalate: 0.2 M oxalic acid) for 16 hours, centrifuging, and then filtering to 0.22 um. Extracts were analyzed for non-crystalline Al and Fe using an ICP.

aige247 - Nutrient Network Oak Litter Decomposition

Decomposition Nutrient Network sites

Decomposition of a common leaf litter substrate was conducted over seven years in the factorial N x P x K+micronutrients experiment and in the factorial fencing x NPK+micronutrients experiment. The decomposition study was carried out at twenty NutNet sites in the United States, Canada, Australia, and Europe, and began less than 1-2 years following the initiation of fertilization treatments. The twenty sites included in the study were: Boulder, Colorado ( Bunchgrass, Oregon ( Bogong High Plains, Victoria ( Burrawan State Forest, NSW ( Chichaqua Bottoms, Iowa ( Cedar Creek, Minnesota ( Cowichan, British Columbia ( Elliott Chaparral Reserve, California ( Halls Prairie, Kentucky ( Hopland Research & Extension Center, California ( Kinypanial, Victoria ( Andrews Lookout, Oregon ( McLaughlin Natural Reserve, California ( Sagehen Creek Field Station, California ( Sedgewick Reserve, California ( Sheep Experiment Station, Idaho ( Sierra Foothill Research & Extension Center, California ( Spindletop Farm, Kentucky ( University of North Carolina ( Val Mustair (


Harvested litter was ground, analyzed for total carbon (C) and N using dry combustion gas chromatography on an Elemental Analyzer (Costech Analytical Technologies Inc., Valencia, CA, USA), and proportion C remaining evaluated for outliers. We analyzed C, rather than mass, remaining to reduce variation associated with soil contamination.

Oak Litter Decomposition litterbags

Leaf litter of Quercus ellipsoidalis was used as a common substrate for decomposition in all sites and treatments in the study. We selected Q. ellipsoidalis litter because it was readily abundant and, since it does not occur in any of the plots, we hoped to avoid home field advantage effects that could confound among-site environmental effects. Litter bags (20 cm x 20 cm, 1-mm mesh) were prepared using freshly fallen leaf litter and deployed between December, 2009 and October, 2010. Sets of seven bags were connected on nylon string and staked individually into plots at the time of deployment. Litter bags were sequentially removed from strings in each plot at approximately annual intervals, and litter was cleaned of any material other than colonizing microbes, dried (65 degrees C to constant mass), weighed, and sent to the University of Minnesota for further processing for total C and N.

ayye247 - Plant above and belowground biomass across an N fertilization gradient after 13 years of fertilization

Biomass measurements - Soil chemistry and carbon stocks - Heterotrophic microbial respiration

Biomass measurements. Aboveground biomass was harvested in two 10 cm x 100 cm strips per plot around peak biomass in mid-August 2020. Biomass from the two strips was pooled, dried to a constant mass at 60 degrees C, weighed to the nearest 0.01g, and scaled to g m-2. The locations of the clipped strips were moved every year to adjacent, but previously unsampled, locations. Belowground biomass (BGB) was harvested in late July 2020 using two 5 cm diameter x 20 cm soil cores per plot. The top 20 cm of soil contains over 80 percent of root biomass at this site. To estimate belowground net primary productivity (BNPP), we removed two 5 cm diameter x 20 cm soil cores from each plot at the beginning of the growing season, and a 1 cm mesh core was placed in the hole. The soil removed from the plot was immediately sieved at 2mm to remove roots, and the sieved soil was placed back into the mesh core in the plot. Cores were placed on May 18, 2020 after the soil had completely thawed and removed on September 13, 2020, prior to the first autumn frost. For both BGB and BNPP, soil was washed off the roots immediately after harvest, and root biomass was dried to a constant mass and weighed to the nearest 0.01g. Root biomass was pooled between cores, and then scaled on an areal basis to g m-2 to compare it directly to aboveground biomass. Soil chemistry and C stocks. To assess the effect of N addition on soil chemistry, we collected 20 cm deep soil cores in each plot in late July 2020. Soil was sieved to 2 mm to remove roots, dried, ground, and weighed before being analyzed for cation exchange capacity (CEC), pH (1:1 v/w in water), and common exchangeable micronutrients using the Mehlich-3 extraction method (Mehlich 1984) (Waypoint Analytical, Tennessee). Soil C and N content were measured on air-dried soils by dry combustion with an ESC 4010 Elemental Analyzer (Costech, Valencia, CA, USA). Soil C stocks were calculated using bulk density measurements taken from microbial respiration soil cores. Heterotrophic microbial respiration. To assess the impact of N addition on heterotrophic microbial respiration, we conducted a 143-day laboratory incubation of soils from each plot, sampled in July 2021. Briefly, 50 g of fresh 2 mm-sieved soil was sampled from the top 10 cm of the mineral soil and adjusted to 70 percent field capacity. Microbial respiration rate (mg C g soil-1 day-1) was measured with a Licor 7000 by sampling the headspace at the beginning and end of a 24-hour period of a sealed 1 L jar containing a soil sample. Between measurements jars with samples were stored in the dark at 20 degrees C and covered with low-density polyethylene film, allowing CO2 and O2 to diffuse freely while retaining water. Respiration was measured for each sample 14 times over the 143-day period. At the beginning of that period, respiration was measured daily to capture the quickly declining decay rate as labile substrates were metabolized; as decay rate slowed as labile substrates were consumed, respiration was measured fortnightly. Cumulative microbial respiration was calculated by averaging microbial respiration rates from consecutive measurements, multiplying by the number of days elapsed between measurements, and adding the previous cumulative measurement.

azye247 - In situ soil respirations throughout the 2020 growing season across an N fertilization gradient

In situ soil respirations

Carbon flux measurements. Soil respiration in field plots was measured biweekly from April through November 2020 (total number of sampling days = 14) using a Licor 6400XT with a soil respiration chamber (Licor, Lincoln, NE). The chamber was fitted onto a 10 cm diameter PVC collar installed at a fixed location in each plot. Three measurements of chamber headspace CO2 concentrations were taken in a fixed location in each plot at each sampling date and then averaged. All aboveground live biomass and litter was cleared from the sampling footprint throughout the season.

azze247 - Net ecosystem exchange measurements throughout the 2020 growing season across an N fertilization gradient

Net Ecosystem Exchange and Gross Primary Productivity

We measured net ecosystem exchange (NEE) by measuring the change in CO2 concentration in the headspace of a chamber (a 1 m3 PVC frame covered with 6 mil clear plastic sheeting with 30 cm flaps that lay on ground) placed over a plot, using a Licor 850-3 (Licor, Lincoln, NE) connected to a Lenovo tablet running the ``Flux Puppy`` software. The chamber was large enough to capture a representative mix of species in each plot, including a dominant perennial bunchgrass (Andropogon gerardii) that often exceeds 0.1 m2 in areal cover. The chamber was sealed to the ground using two heavy chains placed on the ground flaps, and fans were used inside the chamber to ensure air mixing. Light conditions inside the chamber were measured using an Apogee MQ-200X PAR sensor (Apogee Instruments). Chamber CO2 concentrations were measured for at least two minutes or until they stabilized (up to five minutes). To standardize NEE values across light levels, we used garden shade cloth and tarps to create a gradient of decreasing light environments within the chamber. Two-minute measurements were conducted at 50 percent ambient light, 25 percent ambient light, and 0 percent ambient light (i.e. ecosystem respiration, ER). NEE was then calculated using Equation 1: Equation 1: NEE = (p * V/A) * dC/dt Where ??? is the air density (mol air m-3) calculated as P/RT where P is the average atmospheric pressure (Pa), R is the ideal gas constant (8.314 m3 Pa mol air-1 K-1), T is the average temperature in units of Kelvin (K), and dC/dt is the change in CO2-C concentration over time (mol C mol air 1 s-1). The volume of the cube (V) and ground area (A) sampled were 1 m3 and 1 m2, respectively. From the four light measurements, we fit linear and hyperbolic curves to predict NEE at a standard level of photosynthetically active radiation (PAR = 800 micromol m-2 s-1), NEE800, in order to account for variation in ambient light levels during sampling (Kohli et al. 2020). We used the R2 values to assess the fit of each curve, and the hyperbolic estimation was always the better fit. Ecosystem respiration (ER) was estimated using the 0% ambient light measurement. Gross primary productivity (GPP) was estimated using equation 2: Equation 2: GPP = ER - NEE800