The authors acknowledge the support of the Minnesota Corn Research &#38; Promotion Council, the Hueg-Harrison Fellowship, and the Minnesota&#39;s Discovery, Research and InnoVation Economy (MnDRIVE) Fellowship.

1. Field site description
NOTE: When performing 15N tracer field trials, selected sites should minimize variation due to soil, topography, and physical features5. Cross-contamination may occur following lateral soil movement due to slope, wind or water translocation, or tillage while the vertical distribution of soil N may be impacted by subsurface water flow and tile-drainage6.
<ol>
	<li>Describe the experimental field site including past management (e.g., previous crops and tillage), latitude and longitude, soil physical and chemical properties (e.g., soil textural analysis, initial fertility conditions, pH, and soil bulk density).</li>
	<li>Record GPS coordinates for the research site and the field corners.</li>
	<li>Describe growing season management including pest and disease management (herbicide, insecticide, or fungicide use), soil fertility management (including rate, source, placement, and application timing), tillage, irrigation events and amounts, and residue management.</li>
	<li>As crop growth and microbe mediated N transformations are affected by soil moisture, soil temperature, and air temperature, record climate information including daily high and low temperatures, daily precipitation, and soil moisture and temperatures at several depths that reflect the soil sampling depths.</li>
</ol>
2. Plot design
<ol>
	<li>Plant six corn rows (~86,000 plants ha-1) on 76 cm spacing with a final plot dimension of 15.2 m by 4.6 m.
	<ol>
		<li>Establish border areas 1.5 m from each end of the lengthwise dimension (0-1.5 m, 13.7-15.2 m) and an additional border area 1.5 m long (9.8-11.3 m) adjoining the sampling and harvest areas (Figure 1).</li>
		<li>Designate rows 2 and 3 as the in-season plant and soil sampling area (1.5-9.8 m) and rows 4 and 5 as the harvest area (1.5-9.8 m) for corn grain yield.</li>
		<li>Establish a microplot area (11.3-13.7 m) with dimensions of 2.4 m by 3.8 m centered on the width dimension. Collect all 15N enriched plant and soil samples from this area, leaving 0.38 m of unsampled border on the length and width dimensions to minimize edge effects (Figure 2).</li>
	</ol>
	</li>
	<li>Delineate the treatment plot and microplot corners with different colored flags.</li>
</ol>
3. Soil and plant sample precautions
<ol>
	<li>Use dedicated equipment and processing areas for unenriched and enriched materials. Contamination of unenriched materials (fertilizer, soil, or plant) by enriched materials and vice versa can drastically affect results.</li>
	<li>Collect and process 15N enriched soil and plant samples in order of lowest to highest 15N expected enrichment to minimize cross-contamination. Ensure that work surfaces, gloves, utensils, and machinery are thoroughly cleaned between each sample to minimize cross-contamination from sample carryover.</li>
	<li>Minimize foot traffic in microplots to prevent contamination of unenriched sampling areas. Wear protective shoe coverings when accessing microplots and remove them when exiting the microplot area.</li>
</ol>
4. 15N enriched fertilizer preparation and application
<ol>
	<li>Following guidelines put forth by Ref. 2 for fertilizer 15N use efficiency (F15NUE) studies, dilute 10 atom % 15N enriched urea to 5 atom % 15N enriched urea and dissolve in 2 L of deionized water to ensure uniform enrichment of urea fertilizer. 
	NOTE: The required concentration of 15N enriched fertilizer is dependent on the goals of the agronomic study. If the concentration of stock 15N enriched fertilizer exceeds the researcher&#39;s requirements, the stock fertilizer concentration may be diluted with similar conventional fertilizer using the following formula3. 
	X2 = [(C1/C2) - 1] &#215; X1 
	X2 is the mass of the conventional unenriched fertilizer, X1 is the mass of the tracer fertilizer, C1 is the isotopic concentration [expressed as atom % excess (measured atom % enrichment minus the natural background concentration assumed to be 0.3663 atom %)] of the original tracer fertilizer, and C2 is the isotopic concentration of the final mixture. As an example, given 100 g of 10 atom % enriched urea, 92.7 g of conventional unenriched fertilizer would be required for a final isotopic concentration of 5 atom %; 
	X2 = {[(10-0.3663)/5] - 1} &#215; 100.</li>
	<li>Analyze the solution for 15N concentration to verify enrichment. The authors utilized the analytical services provided by UC Davis Stable Isotope Facility. 
	NOTE: Reactions of the soil-plant-microbe regime to fertilizer additions may be affected by the physical form of fertilizer. Depending on the goals of the study, the urea solution may be applied as a liquid or dehydrated to reform crystals. The crystals may be compacted into a cake using a Carver press at 10,000 psi, followed by crushing the cake and screening the particles to the desired size3.</li>
	<li>Evenly apply the 15N enriched urea solutions to the microplots using a calibrated backpack CO2 sprayer (Figure 3A). If multiple N rates or enrichment levels are used, consider using designated CO2 sprayers for each enrichment level or use a single sprayer and apply solutions from the lowest to the highest enrichment to minimize treatment cross-contamination.</li>
	<li>Incorporate urea-containing fertilizers with light tillage, hand rakes, or at least 0.6&#160;cm of irrigation within 24 h of application to minimize volatilization loss potential.</li>
	<li>No additional 15N enriched urea fertilizer is applied to the microplot during the second growing season. Apply conventional unenriched urea to the entire treatment to avoid a differential response in corn growth due to nitrogen.</li>
</ol>
5. Field sample processing: aboveground corn biomass
<ol>
	<li>At each sampling stage, collect a six-aboveground corn plant composite sample from within the sampling area (15N unenriched) and a six-aboveground corn plant composite sample from the 15N enriched microplot. At least two plants should separate each sampled plant to avoid significantly altering plant growth dynamics. The authors collected plant samples at the V8 and R1 corn physiological development stages11 and at physiological maturity (Figure 2).</li>
	<li>Following the principles described in steps 3.1 and 3.2, chop V8 and R1 aboveground biomass (&#8804;5 cm by &#8804;5 cm); a yard waste chipper is a satisfactory option. Place chopped biomass in labeled fabric or paper bags and dry in a forced-air oven at 60 &#176;C until constant mass. Record the biomass dry weight (Figure 3B).</li>
	<li>Partition physiologically mature corn plants into stover (all vegetative tissues including leaves, husks, and stalks), grain, and cob fractions. Chop and dry in a forced-air oven at 60 &#176;C until constant mass. Record the biomass dry weight.</li>
	<li>Within the microplot, cut all corn stalks at the soil surface, tie into a bundle, label according to plot, and remove from the field (Figure 3C). Adjust microplot corner flags to be nearly flush with the soil surface to minimize the risk of removal by the combine during harvest or tillage post-harvest.</li>
	<li>Harvest grain from the harvest area and report yield at 15.5% moisture content12. Harvest remaining research areas with a plot combine.</li>
	<li>Rake unenriched biomass from off the microplot area. Chop and reapply microplot aboveground biomass to the correct plot (Figure 3D).</li>
	<li>Incorporate residue into the soil surface with tillage taking care to minimize soil and corn residue transport into or out of the microplot area. Replace any microplot corner flags removed due to tillage.</li>
	<li>Plant second-year corn on the same rows as the first-year corn.</li>
	<li>Collect second-year aboveground corn biomass only at physiological maturity and process like first-year corn samples as described in step 5.3. Collect microplot samples from the center of the microplot area (1.52 m by 0.76 m) to avoid any potential signal dilution following tillage (Figure 2). Harvest grain from the harvest area and report yield at 15.5% moisture content.</li>
	<li>Following the principles of steps 3.1 and 3.2, thoroughly mix and grind 100 to 200 g of dried plant material to pass through a 2 mm sieve. Thoroughly mix the ground material and store a subsample in a labeled coin envelope for further processing. 
	NOTE: A Thomas Wiley mill is a satisfactory option for plant tissue grinding while a Perten Laboratory Mill 3610 is a satisfactory option for grinding grain. 
	CAUTION: People grinding plant samples should wear ear protection and be protected from inhaling dust by wearing a National Institute for Occupational Safety and Health approved N95 Particulate Filtering Facepiece Respirator.</li>
</ol>
6. Field sample processing: soil
<ol>
	<li>Collect first-year soil samples 8 days after 15N enriched fertilizer application, V8, R1, and post-harvest before tillage. Collect second-year soil samples at pre-plant and post-harvest. Due to logistical sampling constraints, the authors collected in-season soil samples at 0- to 15-, 15- to 30-, and 30- to 60-cm depths, post-harvest soil samples at 0- to 15-, 15- to 30-, 30- to 60-, and 60- to 90-cm depths, and second-year pre-plant soil samples at 0- to 30-, 30- to 60-, 60- to 120- cm depths. 
	NOTE: If a soil probe is unable to collect a soil core to the deepest desired depth as a single core, collect deeper depth cores from the same boreholes as the upper depths discarding the top 1-cm of soil to avoid contamination from soil falling from upper depths.
	<ol>
		<li>Collect a four-core (1.8-cm diameter) composite soil sample from the unenriched sampling area at V8 and R1 using a hand probe. Collect one core in the corn row and three cores between the corn rows.</li>
		<li>Collect a two-core (5-cm diameter) composite soil sample from the unenriched sampling area at pre-plant and post-harvest using a hydraulic probe.</li>
		<li>Collect a 15-core (1.8-cm diameter) composite soil sample from the microplot area 8 days after 15N enriched fertilizer application, V8, and R1 using a hand probe. Collect three to four cores in the corn row and 11 to 12 cores between the corn rows. 
		NOTE: Soils are extremely heterogeneous. The greater number of cores collected from within the enriched microplot provides a better estimate of the true 15N enrichment of soil N13.</li>
		<li>Collect a three-core (5-cm diameter) composite soil sample from the microplot area at pre-plant and post-harvest using a hydraulic probe.</li>
		<li>Homogenize each composite soil sample in a bucket and place it in a pre-labeled paper bag.</li>
	</ol>
	</li>
	<li>Dry soil samples at 35 &#176;C in a forced-air oven until constant mass. Grind each sample to pass through a 2 mm sieve. A mechanical soil grinder is satisfactory if it can be thoroughly cleaned between each sample. 
	NOTE: Soil samples may be air-dried by spreading samples on trays in a thin layer. Trays should be in an area free from contamination by outside N sources. Unenriched and enriched samples should be physically separated to prevent cross-contamination. 
	CAUTION: People grinding soil samples should wear ear protection and be protected from inhaling dust by wearing a National Institute for Occupational Safety and Health approved N95 Particulate Filtering Facepiece Respirator.</li>
</ol>
7. Lab sample processing: grind soil and plant samples
<ol>
	<li>Dry ground plant samples (2 mm) overnight in an oven at 60 &#176;C.</li>
	<li>Following the principles described in step 3, grind dried plant samples or soil material to a fine, flour-like consistency. A roller jar mill is a satisfactory option. 
	NOTE: The authors&#39; jar mill is a custom-built conveyor belt system that can process 54 roller jars at a time.
	<ol>
		<li>Fill each roller jar (250 mL borosilicate glass jar with a screw-top lid) with 10 to 20 g of ground plant or soil sample and seven stainless steel rods (8.5 cm long, 0.7 cm diameter).</li>
		<li>Roll roller jars at 0.4 x g for 6-24 h or until samples have a fine, flour-like consistency.</li>
		<li>Transfer the finely ground material into a clean, labeled 20 mL scintillation vial.</li>
		<li>Between each sample, wash roller jars, stainless steel rods, and lids with soap and water to remove any residue.
		<ol>
			<li>Immerse roller jars and lids in a 5% HCl acid bath (prepared from 36-38% concentrated stock) overnight14. 
			CAUTION: Hydrochloric acid is corrosive. It can cause severe skin burns, eye damage, and is harmful if inhaled. Always wear protective clothing, gloves, and eye and face protection. Flush contacted tissue thoroughly with water. Always use a secondary container when transporting acids. Always add acid to water as this reaction is exothermic. Immediately neutralize acid spills with baking soda. 
			NOTE: A large acid bath may be prepared as 100 L of 5% HCl in a 208 L plastic container. Prepare several smaller volumes in a fume hood and then transfer the solutions to the plastic container. Replace the solution every three months.</li>
			<li>Triple rinse roller jars and lids with deionized water and air dry.</li>
			<li>Immerse stainless steel rods in a 0.05 M NaOH bath (prepared by dissolving 2 g of NaOH in 1 L of deionized water) overnight14. Prepare a new 0.05 M NaOH bath each day. 
			CAUTION: Sodium hydroxide can cause severe skin burns and eye damage. Always wear protective clothing and eye protection. Immediately remove contaminated clothing and rinse skin or eyes with water for several minutes.</li>
			<li>Rinse the rods under running hot tap water for 5 minutes. Decant and triple rinse the rods with deionized water. Allow the rods to air dry on a paper towel-lined tray.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
8. Weigh ground plant and soil samples for total N and 15N analysis
<ol>
	<li>Analyze a few representative plant and soil samples for total N content (e.g., combustion analysis15). Calculate the sample mass that provides adequate N content for 15N analysis according to the analyzer specifications. 
	NOTE: The authors utilized the analytical services provided by UC Davis Stable Isotope Facility. Enriched sample weights were optimized for 20 &#181;g of N with a maximum of 100 &#181;g of N.</li>
	<li>Organize like-samples from lowest to highest expected 15N enrichment. Duplicate every eighth to twelfth sample in each run to check sample precision. Include at least one check sample per run16.</li>
	<li>Label a clean 96-well plate and fitted lid with individual well evaporation rings. Cut a clean index card to fit just inside the lid to prevent sample movement between wells during transport.</li>
	<li>Wearing nitrile gloves, clean the microscale, work surfaces, spatula, and forceps with laboratory wipes and ethanol. Place cleaned utensils on a Kimwipe on the lab bench. 
	NOTE: Unenriched and enriched samples should be processed using separate scales and utensils to prevent cross-contamination.</li>
	<li>Use forceps to place a pre-formed 5 mm x 9 mm tin capsule on a clean work surface, such as a stainless steel block with 5 mm x 8 mm well. Gently tap the capsule into the well to reform the cylindrical shape and flatten the bottom of the capsule if needed. 
	NOTE: Because sample masses will be very small, the risk of sample contamination is high. Never touch the capsules with gloves. Discard the capsule if it touches any surface other than the forceps, clean work surface, scale weigh pan, or 96-well plate.</li>
	<li>Use forceps to gently flare out the top 1 mm of the capsule to facilitate manipulation. To avoid scale damage when taring the weight of the capsule, hover and release the capsule 1 to 2 mm above the microscale weigh pan. Tare the capsule. Use forceps to return the capsule to the clean work surface.</li>
	<li>Use a spatula to carefully add the required mass of finely ground sample material to the capsule. Avoid spilling sample material on the outside surface of the capsule or the work surfaces.</li>
	<li>Using forceps, slowly crimp the top third of the capsule and fold over to seal. Using forceps, continue to fold and compress the capsule into a spherical shape taking care not to puncture or tear the tin. 
	NOTE: Samples with low N content may require sample volumes that exceed the capacity of the 5x9 mm capsule. Larger capsules (e.g., 9 mm x 10 mm) may be used in these instances.</li>
	<li>Use forceps to drop the wrapped capsule several times from a height of 1 cm onto a clean, dark surface or mirror to check for leaks. If no dust appears, weigh the sample using the same technique as described in step 8.6. Record the sample weight. Place the capsule in a 96-well plate and record the well placement.
	<ol>
		<li>If dust appears on the dark surface, record the sample weight. Wrap the sample in a second tin capsule, recheck for leaks, and place it in a clean 96-well plate. 
		NOTE: If the wrapped capsule is too large to fit in a 96-well plate, use a 24- or 48-well plate.</li>
	</ol>
	</li>
	<li>Between samples, clean each of the utensils and surfaces with ethanol and laboratory wipes paying especial attention to the spatula and forceps edges.</li>
	<li>Secure the lid to the 96-well plate using tape and store in a desiccator.</li>
</ol>
9. Calculations
<ol>
	<li>Calculate the mass of N (kg&#8729;ha-1) contained in the plant or soil samples using the following equations. 
	<img src="/files/ftp_upload/61191/61191eq1.jpg" /> 
	<img src="/files/ftp_upload/61191/61191eq2.jpg" /></li>
	<li>Calculate the fertilizer N fraction (Nf), fertilizer derived N (FDN), and soil derived N (SDN) for plant and soil samples17. 
	<img src="/files/ftp_upload/61191/61191eq3.jpg" /> 
	where A is the atom % 15N enrichment.</li>
	<li><img src="/files/ftp_upload/61191/61191eq4.jpg" /> 
	<img src="/files/ftp_upload/61191/61191eq5.jpg" /></li>
	<li>Calculate fertilizer 15N use efficiency17. 
	<img src="/files/ftp_upload/61191/61191eq6v2.jpg" /></li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fate+of15N-labelled+fertilizer+applied+to+spring+barley+grown+on+soils+of+contrasting+nutrient+status.">Fate of15N-labelled fertilizer applied to spring barley grown on soils of contrasting nutrient status.</a> Plant and Soil. 195 (1), 83-98 (1997).</li><li>Khanif, Y. M., Cleemput, O., Baert, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Field+study+of+the+fate+of+labelled+fertilizer+nitrate+applied+to+barley+and+maize+in+sandy+soils.">Field study of the fate of labelled fertilizer nitrate applied to barley and maize in sandy soils.</a> Fertilizer Research. 5 (3), 289-294 (1984).</li></ol>

The authors have nothing to disclose.

Stable isotope research is a useful tool for tracking and quantifying FDN through the soil-crop system. However, there are three main assumptions associated with N tracer studies that if violated may invalidate conclusions drawn from using this methodology. They are 1) the tracer is uniformly distributed throughout the system, 2) processes under the study occur at the same rates, and 3) N leaving the 15N enriched pool does not return3. Because this study is interested in the distributio...

Fertilizer nitrogen (N) use is essential in agriculture to meet the food, fiber, feed, and fuel demands of a growing global population, but N losses from agricultural fields can negatively impact environmental quality. Because N undergoes many transformations in the soil-crop system, a better understanding of N cycling, crop utilization, and the overall fate of fertilizer N are necessary to improve management practices that promote N use efficiency and minimize environmental losses. Traditional N fertilizer studies primarily focus on the effect of a treatment on end-of-season measurements such as crop yield, crop N uptake relative to the N rate applied (apparent fertilizer use efficiency), and residual soil N. While these studies quantify the overall system N inputs, outputs, and efficiencies, they cannot identify nor quantify N in the soil-crop system derived from fertilizer sources or the soil. A different approach using stable isotopes must be used to track and quantify the fate of fertilizer derived N (FDN) in the soil-crop system.
Nitrogen has two stable isotopes, 14N and 15N, that occur in nature at a relatively constant ratio of 272:1 for 14N/15N1 (concentration of 0.366 atom % 15N or 3600 ppm 15N2,3). The addition of 15N enriched fertilizer increases the total 15N content of the soil system. As 15N enriched fertilizer mixes with unenriched soil N, the measured change of 14N/15N ratio allows researchers to trace FDN in the soil profile and into the crop3,4. A mass balance can be calculated by measuring the total amount of 15N tracer in the system and each of its parts2. Because 15N enriched fertilizers are significantly more expensive than conventional fertilizers, 15N enriched microplots are often embedded within the treatment plots. The purpose of this methods paper is to describe a small-plot research&#160;design utilizing microplots for multiple in-season soil and plant sampling events for corn (Zea mays L.) and to present protocols for preparing plant and soil samples for total 15N analysis. These results can then be used to estimate N fertilizer use efficiency and create a partial N budget accounting for FDN in the bulk soil and the crop.

<table><tbody><tr><td>20 mL scintillation vial</td><td>ANY; Fisher Scientific is one example</td><td>0334172C</td><td></td></tr><tr><td>250 mL borosilicate glass bottle</td><td>QORPAK</td><td>264047</td><td></td></tr><tr><td>48-well plate</td><td>EA Consumables</td><td>E2063</td><td></td></tr><tr><td>96-well plate</td><td>EA Consumables</td><td>E2079</td><td></td></tr><tr><td>Cloth parts bag (30x50 cm)</td><td>ANY</td><td>NA</td><td>For corn ears</td></tr><tr><td>CO2 Backpack Sprayer</td><td>ANY; Bellspray Inc is one example</td><td>Model T</td><td></td></tr><tr><td>Coin envelop (6.4x10.8 cm)</td><td>ANY; ULINE is one example</td><td>S-6285</td><td>For 2-mm ground plant samples</td></tr><tr><td>Corn chipper</td><td>ANY; DR Chipper Shredder is one example</td><td>SKU:CS23030BMN0</td><td>For chipping corn biomass</td></tr><tr><td>Corn seed</td><td>ANY</td><td>NA</td><td>Hybrid appropriate to the region</td></tr><tr><td>Disposable shoe cover</td><td>ANY; Boardwalk is one example</td><td>BWK00031L</td><td></td></tr><tr><td>Ethanol 200 Proof</td><td>ANY; Decon Laboratories Inc. is one example</td><td>2701TP</td><td></td></tr><tr><td>Fabric bags with drawstring (90x60 cm)</td><td>ANY</td><td>NA</td><td>For plant sample collection</td></tr><tr><td>Fertilizer Urea (46-0-0)</td><td>ANY</td><td>NA</td><td>~0.366 atom % 15N</td></tr><tr><td>Hand rake</td><td>ANY; Fastenal Company is one example</td><td>5098-63-107</td><td></td></tr><tr><td>Hand sickle</td><td>ANY; Home Depot is one example</td><td>NJP150</td><td>For plant sample collection</td></tr><tr><td>Hand-held soil probe</td><td>ANY; AMS is one example</td><td>401.01</td><td></td></tr><tr><td>Hydraulic soil probe</td><td>ANY; Giddings is one example</td><td>GSPS</td><td></td></tr><tr><td>Hydrochloric acid, 12N</td><td>Ricca Chemical</td><td>R37800001A</td><td></td></tr><tr><td>Jar mill</td><td>ANY; Cole-Parmer is one example</td><td>SI-04172-50</td><td></td></tr><tr><td>Laboratory Mill</td><td>Perten</td><td>3610</td><td>For grinding grain</td></tr><tr><td>Microbalance accurate to four decimal places</td><td>ANY; Mettler Toledo is one example</td><td>XPR2</td><td></td></tr><tr><td>N95 Particulate Filtering Facepiece Respirator</td><td>ANY, ULINE is one example</td><td>S-9632</td><td></td></tr><tr><td>Neoprene or butyl rubber gloves</td><td>ANY</td><td>NA</td><td>For working in HCl acid bath</td></tr><tr><td>Paper hardware bags (13.3x8.7x27.8 cm)</td><td>ANY; ULINE is one example</td><td>S-8530</td><td>For soil samples and corn grain</td></tr><tr><td>Plant grinder</td><td>ANY; Thomas Wiley Model 4 Mill is one example</td><td>1188Y47-TS</td><td>For grinding chipped corn biomass to 2-mm particles</td></tr><tr><td>Plastic tags</td><td>ULINE</td><td>S-5544Y-PW</td><td>For labeling fabric bags and microplot stalk bundles</td></tr><tr><td>Sodium hydroxide pellets, ACS</td><td>Spectrum Chemical</td><td>SPCM-S1295-07</td><td></td></tr><tr><td>Soil grinder</td><td>ANY; AGVISE stainless steel grinder with motor is one example</td><td>NA</td><td>For grinding soil to pass through a 2-mm sieve</td></tr><tr><td>Tin capsule 5x9 mm</td><td>Costech Analytical Technologies Inc.</td><td>041061</td><td></td></tr><tr><td>Tin capsule 9x10 mm</td><td>Costech Analytical Technologies Inc.</td><td>041073</td><td></td></tr><tr><td>Urea (46-0-0)</td><td>MilliporeSigma</td><td>490970</td><td>10 atom % 15N</td></tr></tbody></table>

microplot design and plant and soil sample preparation for 15nitrogen analysis

Many&#160;nitrogen fertilizer studies evaluate the overall effect of a treatment on end-of-season measurements such as grain yield or cumulative N losses. A stable isotope approach is necessary to follow and quantify the fate of fertilizer derived N (FDN) through the soil-crop system. The purpose of this paper is to describe a small-plot research&#160;design utilizing non-confined 15N enriched microplots for multiple soil and plant sampling events over two growing seasons and provide sample collection, handling, and processing protocols for total 15N analysis. The methods were demonstrated using a replicated study from south-central Minnesota planted to corn (Zea mays L.). Each treatment consisted of six corn rows (76 cm row-spacing) 15.2 m long with a microplot (2.4 m by 3.8 m) embedded at one end. Fertilizer-grade urea was applied at 135 kg N&#8729;ha-1 at planting, while the microplot received urea enriched to 5 atom % 15N. Soil and plant samples were taken several times throughout the growing season, taking care to minimize cross-contamination by using separate tools and physically separating unenriched and enriched samples during all procedures. Soil and plant samples were dried, ground to pass through a 2 mm screen, and then ground to a flour-like consistency using a roller jar mill. Tracer studies require additional planning, sample processing time and manual labor, and incur higher costs for 15N enriched materials and sample analysis than traditional N studies. However, using the mass balance approach, tracer studies with multiple in-season sampling events allow the researcher to estimate FDN distribution through the soil-crop system and estimate unaccounted-for FDN from the system.

The results presented in this paper come from a field site established in 2015 at the University of Minnesota Southern Outreach and Research Center located near Waseca, MN. The site was managed as a corn-soybean [Glycine max (L.) Merr] rotation prior to 2015 but was managed as a corn-corn rotation during the 2015 and 2016 growing seasons. The soil was a Nicollet clay loam (fine-loamy, mixed, superactive, mesic Aquic Hapludolls)-Webster clay loam (fine-loamy, mixed, superactive, m...

Watch this Scientific Journal Video about Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis at JoVE.com

Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis

A microplot design for 15N tracer research is described to accommodate multiple in-season plant and soil sampling events. Soil and plant sample collection and processing procedures, including grinding and weighing protocols, for 15N analysis are put forth.

Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis

Many&#160;nitrogen fertilizer studies evaluate the overall effect of a treatment on end-of-season measurements such as grain yield or cumulative N losses. ...

microplot-design-plant-soil-sample-preparation-for-15nitrogen

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6.7K Views.  University of Minnesota. A microplot design for 15N tracer research is described to accommodate multiple in-season plant and soil sampling events. Soil and plant sample collection and processing procedures, including grinding and weighing protocols, for 15N analysis are put forth.

Video: Microplot Design and Plant and Soil Sample Preparation for 15Nitrogen Analysis

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes and soil organic matter formation to small-scale stable-isotope biomarker probing. Coupling multiple stable isotopes such as 13C with 15N, 18O or&#160;2H has the potential to reveal even more information about complex stoichiometric relationships during biogeochemical transformations. Isotope labeled plant material has been used in various studies of litter decomposition and soil organic matter formation1-4. From these and other studies, however, it has become apparent that structural components of plant material behave differently than metabolic components (i.e. leachable low molecular weight compounds) in terms of microbial utilization and long-term carbon storage5-7. The ability to study structural and metabolic components separately provides a powerful new tool for advancing the forefront of ecosystem biogeochemical studies. Here we describe a method for producing 13C and 15N labeled plant material that is either uniformly labeled throughout the plant or differentially labeled in structural and metabolic plant components.
Here, we present the construction and operation of a continuous 13C and 15N labeling chamber that can be modified to meet various research needs. Uniformly labeled plant material is produced by continuous labeling from seedling to harvest, while differential labeling is achieved by removing the growing plants from the chamber weeks prior to harvest. Representative results from growing Andropogon gerardii Kaw demonstrate the system&#39;s ability to efficiently label plant material at the targeted levels. Through this method we have produced plant material with a 4.4 atom%13C and 6.7 atom%15N uniform plant label, or material that is differentially labeled by up to 1.29 atom%13C and 0.56 atom%15N in its metabolic and structural components (hot water extractable and hot water residual components, respectively). Challenges lie in maintaining proper temperature, humidity, CO2 concentration,&#160;and light levels in an airtight 13C-CO2 atmosphere for successful plant production. This chamber description represents a useful research tool to effectively produce uniformly or differentially multi-isotope labeled plant material for use in experiments on ecosystem biogeochemical cycling.

Design and Operation of a Continuous 13C and 15N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

This method explains how to build and operate a continuous 13C and 15N isotope labeling chamber for uniform or differential plant tissue labeling. Representative results from metabolic and structural labeling of Andropogon gerardii are discussed.

Tracing rare stable isotopes from plant material through the ecosystem provides the most sensitive information about ecosystem processes; from CO2 fluxes ...

Calibrated Passive Sampling - Multi-plot Field Measurements of NH3 Emissions with a Combination of Dynamic Tube Method and Passive Samplers

Agricultural ammonia (NH3) emissions (90% of total EU emissions) are responsible for about 45% airborne eutrophication, 31% soil acidification and 12% fine dust formation within the EU15. But NH3 emissions also mean a considerable loss of nutrients. Many studies on NH3 emission from organic and mineral fertilizer application have been performed in recent decades. Nevertheless, research related to NH3 emissions after application fertilizers is still limited in particular with respect to relationships to emissions, fertilizer type, site conditions and crop growth. Due to the variable response of crops to treatments, effects can only be validated in experimental designs including field replication for statistical testing. The dominating ammonia loss methods yielding quantitative emissions require large field areas, expensive equipment or current supply, which restricts their application in replicated field trials. This protocol describes a new methodology for the measurement of NH3 emissions on many plots linking a simple semi-quantitative measuring method used in all plots, with a quantitative method by simultaneous measurements using both methods on selected plots. As a semi-quantitative measurement method passive samplers are used. The second method is a dynamic chamber method (Dynamic Tube Method) to obtain a transfer quotient, which converts the semi-quantitative losses of the passive sampler to quantitative losses (kg nitrogen ha-1). The principle underlying this approach is that passive samplers placed in a homogeneous experimental field have the same NH3 absorption behavior under identical environmental conditions. Therefore, a transfer co-efficient obtained from single passive samplers can be used to scale the values of all passive samplers used in the same field trial. The method proved valid under a wide range of experimental conditions and is recommended to be used under conditions with bare soil or small canopies (&#60;0.3 m). Results obtained from experiments with taller plants should be treated more carefully.

Calibrated Passive Sampling - Multi-plot Field Measurements of NH3 Emissions with a Combination of Dynamic Tube Method and Passive Samplers

Ammonia emissions are a major threat to the environment by eutrophication, soil acidification and fine particle formation and stem mainly from agricultural sources. This method allows ammonia loss measurements in replicated field trials enabling statistical analysis of emissions and of relationships between crop development and emissions.

Agricultural ammonia (NH3) emissions (90% of total EU emissions) are responsible for about 45% airborne eutrophication, 31% soil acidification and 12% ...

A Protocol for Collecting and Constructing Soil Core Lysimeters

Leaching of nutrients from land applied fertilizers and manure used in agriculture can lead to accelerated eutrophication of surface water. Because the landscape has complex and varied soil morphology, an accompanying disparity in flow paths for leachate through the soil macropore and matrix structure is present. The rate of flow through these paths is further affected by antecedent soil moisture. Lysimeters are used to quantify flow rate, volume of water and concentration of nutrients leaching downward through soils. While many lysimeter designs exist, accurately determining the volume of water and mass balance of nutrients is best accomplished with bounded lysimeters that leave the natural soil structure intact. 
Here we present a detailed method for the extraction and construction of soil core lysimeters equipped with soil moisture sensors at 5 cm and 25 cm depths. Lysimeters from four different Coastal Plain soils (Bojac, Evesboro, Quindocqua and Sassafras) were collected on the Delmarva Peninsula and moved to an indoor climate controlled facility. Soils were irrigated once weekly with the equivalent of 2 cm of rainfall to draw down soil nitrate-N concentrations. At the end of the draw down period, poultry litter was applied (162 kg TN ha-1) and leaching was resumed for an additional five weeks. Total recovery of applied irrigation water varied from 71% to 85%. Nitrate-N concentration varied over the course of the study from an average of 27.1 mg L-1 before litter application to 40.3 mg L-1 following litter application. While greatest flux of nutrients was measured in soils dominated by coarse sand (Sassafras) the greatest immediate flux occurred from the finest textured soil with pronounced macropore development (Quindocqua).

A detailed method for extraction and assembly of intact soil core lysimeters and their use for study of leachate and associated loss of nutrients from surface applied poultry litter is demonstrated.

Leaching of nutrients from land applied fertilizers and manure used in agriculture can lead to accelerated eutrophication of surface water. Because the ...

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Nitrogen oxides (NOx = NO + NO2) are a family of atmospheric trace gases that have great impact on the environment. NOx concentrations directly influence the oxidizing capacity of the atmosphere through interactions with ozone and hydroxyl radicals. The main sink of NOx is the formation and deposition of nitric acid, a component of acid rain and a bioavailable nutrient. NOx is emitted from a mixture of natural and anthropogenic sources, which vary in space and time. The collocation of multiple sources and the short lifetime of NOx make it challenging to quantitatively constrain the influence of different emission sources and their impacts on the environment. Nitrogen isotopes of NOx have been suggested to vary amongst different sources, representing a potentially powerful tool to understand the sources and transport of NOx. However, previous methods of collecting atmospheric NOx integrate over long (week to month) time spans and are not validated for the efficient collection of NOx in relevant, diverse field conditions. We report on a new, highly efficient field-based system that collects atmospheric NOx for isotope analysis at a time resolution between 30 min and 2 hr. This method collects gaseous NOx in solution as nitrate with 100% efficiency under a variety of conditions. Protocols are presented for collecting air in urban settings under both stationary and mobile conditions. We detail the advantages and limitations of the method and demonstrate its application in the field. Data from several deployments are shown to 1) evaluate field-based collection efficiency by comparisons with in situ NOx concentration measurements, 2) test the stability of stored solutions before processing, 3) quantify in situ reproducibility in a variety of urban settings, and 4) demonstrate the range of N isotopes of NOx detected in ambient urban air and on heavily traveled roadways.

Automated, High-resolution Mobile Collection System for the Nitrogen Isotopic Analysis of NOx

Previous work suggests that the nitrogen isotopic composition of atmospheric nitrogen oxides might distinguish the influence of different sources in the environment. We report on an automated, mobile, field-based method for the high collection efficiency of atmospheric NOx for N isotopic analysis at an hourly time resolution.

Nitrogen oxides (NOx = NO + NO2) are a family of atmospheric trace gases that have great impact on the environment. NOx concentrations directly influence ...

Isolation and Analysis of Microbial Communities in Soil, Rhizosphere, and Roots in Perennial Grass Experiments

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The purpose of this manuscript is to provide detail on how to rapidly sample soil, rhizosphere, and endosphere of replicated field trials and analyze changes that may occur in the microbial communities due to sample type, treatment, and plant genotype. The experiment used to demonstrate these methods consists of replicated field plots containing two, pure, warm-season grasses (Panicum virgatum and Andropogon gerardii) and a low-diversity grass mixture (A. gerardii, Sorghastrum nutans, and Bouteloua curtipendula). Briefly, plants are excavated, a variety of roots are cut and placed in phosphate buffer, and then shaken to collect the rhizosphere. Roots are brought to the laboratory on ice and surface sterilized with bleach and ethanol (EtOH). The rhizosphere is filtered and concentrated by centrifugation. Excavated soil from around the root ball is placed into plastic bags and brought to the lab where a small amount of soil is taken for DNA extractions. DNA is extracted from roots, soil, and rhizosphere and then amplified with primers for the V4 region of the 16S rRNA gene. Amplicons are sequenced, then analyzed with open access bioinformatics tools. These methods allow researchers to test how the microbial community diversity and composition varies due to sample type, treatment, and plant genotype. Using these methods along with statistical models, the representative results demonstrate there are significant differences in microbial communities of roots, rhizosphere, and soil. Methods presented here provide a complete set of steps for how to collect field samples, isolate, extract, quantify, amplify, and sequence DNA, and analyze microbial community diversity and composition in replicated field trials.

Excavation of plant roots from the field as well as processing of samples into endosphere, rhizosphere, and soil are described in detail, including DNA extraction and data analysis methods. This paper is designed to enable other laboratories to use these techniques for the study of soil, endosphere, and rhizosphere microbiomes.

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The ...

Sampling Soils in a Heterogeneous Research Plot

Soils are highly heterogeneous. In general, the number of soil samples required for soil research has always been determined arbitrarily and the associated accuracy is unknown. Here, we present a detailed protocol for efficient and clustered soil sampling in a research plot and, relying on a pilot sampling using this design, for demonstrating soil spatial heterogeneity and informing reasonable sample sizes and associated accuracy for future study. The protocol mainly comprises four steps: sampling design, field collection, soil analysis, and geostatistical analysis. The step-by-step procedure is modified according to former publications. Two examples will be presented to demonstrate contrasting spatial distributions of soil organic carbon (SOC) and soil microbial biomass carbon (MBC) under different management practices. In addition, we present a strategy to determine the sample size requirement (SSR) given a certain level of accuracy based on the plot-level coefficient of variation (CV). The field sampling protocol and the quantitative determination of the sample size will assist researchers in seeking feasible sampling strategies to meet research needs and resources' availability.

The traditional soil-sampling procedure determines the number of soil samples arbitrarily. Here, we provide a simple yet efficient clustered soil-sampling design to demonstrate soil spatial heterogeneity and quantitatively determine the number of soil samples required and the associated sampling accuracy.

Soils are highly heterogeneous. In general, the number of soil samples required for soil research has always been determined arbitrarily and the ...

An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive harvest or expensive equipment. We present a customizable and affordable rhizobox method that allows the non-destructive visualization of root growth over time and is particularly well-suited to studying root plasticity in response to localized resource patches. The method was validated by assessing maize genotypic variation in plasticity responses to patches containing 15N-labeled legume residue. Methods are described to obtain representative developmental measurements over time, measure root length density in resource-containing and control patches, calculate root growth rates, and determine 15N recovery by plant roots and shoots. Advantages, caveats, and potential future applications of the method are also discussed. Although care must be taken to ensure that experimental conditions do not bias root growth data, the rhizobox protocol presented here yields reliable results if carried out with sufficient attention to detail.

Visualizing and measuring root growth in situ is extremely challenging. We present a customizable rhizobox method to track root development and proliferation over time in response to nutrient enrichment. This method is used to analyze maize genotypic differences in root plasticity in response to an organic nitrogen source.

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive ...

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has been increasing because global nitrogen loads have dramatically increased following industrialization. Dissimilatory nitrate reduction to ammonium (DNRA) and denitrification are both microbial processes that use NO3&#8722; for respiration. Compared to denitrification, quantitative determinations of the DNRA activity have been carried out only to a limited extent. This has led to an insufficient understanding of the importance of DNRA in NO3&#8722; transformations and the regulating factors of this process. The objective of this paper is to provide a detailed procedure for the measurement of the potential DNRA rate in environmental samples. In brief, the potential DNRA rate can be calculated from the 15N-labeled ammonium (15NH4+) accumulation rate in 15NO3&#8722; added incubation. The determination of the 14NH4+ and 15NH4+ concentrations described in this paper is comprised of the following steps. First, the NH4+ in the sample is extracted and trapped on an acidified glass filter as ammonium salt. Second, the trapped ammonium is eluted and oxidized to NO3&#8722; via persulfate oxidation. Third, the NO3&#8722; is converted to N2O via an N2O reductase deficient denitrifier. Finally, the converted N2O is analyzed using a previously developed quadrupole gas chromatography&#8211;mass spectrometry system. We applied this method to salt marsh sediments and calculated their potential DNRA rates, demonstrating that the proposed procedures allow a simple and more rapid determination compared to previously described methods.

Measurement of the Potential Rates of Dissimilatory Nitrate Reduction to Ammonium Based on 14NH4+/15NH4+ Analyses via Sequential Conversion to N2O

A series of methods to determine the potential DNRA rate based on 14NH4+/15NH4+ analyses is provided in detail. NH4+ is converted into N2O via several steps and analyzed using quadrupole gas chromatography&#8211;mass spectrometry.

The importance of understanding the fate of nitrate (NO3&#8722;), which is the dominant N species transferred from terrestrial to aquatic ecosystems, has ...

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

Carbon and Nitrogen Analysis of Environmental Samples

Source: Laboratories of Margaret Workman and Kimberly Frye - Depaul University
Elemental Analysis is a method used to determine elemental composition of a material. In environmental samples such as soils, scientists are particularly interested in the amounts of two ecologically important elements, nitrogen and carbon. Elemental analysis by the flash combustion technique works by oxidizing the sample with a catalyst through combustion in a high-temperature chamber. The products of combustion are then reduced to N2 and CO2 and detected with a thermal conductivity detector.
Unlike other methods for total nitrogen determination (Kjeldahl method) and total carbon determination (Walkley-Black, Heanes or Leco methods), the flash combustion technique does not use toxic chemicals and is therefore much safer to use.
This video will demonstrate combustion-based elemental analysis using the Flash EA 1112 instrument from Thermo Fisher Scientific.

Source: Laboratories of Margaret Workman and Kimberly Frye - Depaul University
Elemental Analysis is a method used to determine elemental composition of a ...

Microplot Design and Plant and Soil Sample Preparation for ¹⁵Nitrogen Analysis

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Chapters in this video

Design and Operation of a Continuous ¹³C and ¹⁵N Labeling Chamber for Uniform or Differential, Metabolic and Structural, Plant Isotope Labeling

Calibrated Passive Sampling - Multi-plot Field Measurements of NH₃ Emissions with a Combination of Dynamic Tube Method and Passive Samplers