The authors gratefully acknowledge financial support from The Scotts Miracle-Gro Company for this facility. We are also appreciative to the Toro Co. for assistance with providing the irrigation controller. The vision and planning by the late Dr. Chris Steigler in the early stages of this project is also gratefully acknowledged. The authors would also like to thank Ms. N. Stanley for her technical assistance with sample preparation and analysis.

1. Site Selection
<ol>
	<li>Locate a suitably-sized area of undisturbed soil having a uniform 3-4% slope.</li>
	<li>Conduct a topographic survey and delineate an area approximately 10 m x&#160;100 m having an average 3.7&#177;0.5% slope.</li>
	<li>Divide the 10 m x&#160;100 m area into three blocks, each approximately 10 m x&#160;33.3 m (Figure 1).</li>
	<li>Subdivide each block into 8 field plots, each 4.1 m wide by 8.2 m long.</li>
	<li>Identify and document the soil series present in the study area. Note: this location had a Booneville series fine sandy loam but other soil series and textures may be used.</li>
</ol>
2. Retaining Wall Construction
<ol>
	<li>Cut a 30 cm wide by 30 cm deep trench at the low end of the plots.</li>
	<li>Cut a 20 cm wide by 1.2 m deep trench 10 cm from the plot edge to provide a smooth vertical edge that extends into the clay subsoil.</li>
	<li>Construct and install temporary wooden forms in the trench to hold it open.</li>
	<li>Remove soil adjacent to the downhill side of the form to a depth of 76 cm below the soil surface at the low end of the plots. Insure a minimum slope of 0.5% away from the plots for a distance of approximately 30 m to provide adequate drainage.</li>
	<li>Remove the temporary forms and construct a steel reinforced concrete retaining wall.
	<ol>
		<li>Construct wooden forms for the outside of the wall and use the undisturbed soil below the plot areas as the inside wall.</li>
		<li>Be sure that the wall extends into undisturbed subsoil to help prevent future movement.</li>
		<li>Assemble two sections of trench drain for each plot with end caps at each end and a bottom discharge drain at the low end. Seal all joints with silicone and then screw the joints together as per manufacturer&#8217;s recommendations.</li>
		<li>Glue and screw a 10 cm diameter PVC 90&#176; ell and 60 cm length of discharge pipe to the outlet. Place the assembled drain in the concrete form and attach it so that the top edge is level in both directions and 1.27 cm below the soil surface at the low end of the plot (Figure 2). Cover the drain with a temporary plastic cover to keep out wet concrete.</li>
		<li>Pour 4,000 lb. test ready mix concrete into the forms using appropriate amounts of vibration to remove voids.
		<ol>
			<li>When forms are full, trowel the upper surface to form a smooth finish with rounded edges. Temporary plastic covers on the drains should be removed to allow final surface preparation.</li>
			<li>Ensure that the finished concrete surface is level with the soil surface at the bottom of the plot and has a 1.27 cm slope to the drain.</li>
			<li>Ensure that, on the downhill side of the drain, the concrete has a 1.27 cm slope away from the drain to prevent water from backing up into the drains.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Form and pour steel reinforced concrete pads (1.2 m wide, 1.8 m long, and 15 cm thick) below each drain outflow. Pads must have 0.5% slope away from the wall and the top of the pad must be 30 cm below the bottom of the drain outlet.</li>
	<li>Provide a weatherproof electrical outlet (110/120 V) on the side of the retaining wall above each pad in preparation for instrumentation.</li>
</ol>
3. Installation of Instrumentation
<ol>
	<li>Cut discharge pipes flush with the concrete wall.</li>
	<li>Install a 1.2 m long H flume immediately below the drain outflow.
	<ol>
		<li>Anchor the flume to the wall using appropriate concrete anchors and screws being sure that the flume is level from side to side.</li>
		<li>Support the front of the flume with an adjustable stainless steel stand and use the adjustments to level the unit both side to side and front to back. Seal the joints between the flumes and concrete with Tub and Tile sealant.</li>
	</ol>
	</li>
	<li>Install a flow meter on each pad. Locate the flow meter near the end of the flume to minimize the length of tubing needed.</li>
	<li>Install a portable sampler on each pad. Locate the sampler as needed to minimize the needed amount of tubing to reach the sampling tube. Note: It may be necessary to put the sampler on a stand to prevent depressions that may retain water in the sampling tubing.</li>
	<li>Design, fabricate, and install stainless steel covers over the wall and the flumes to prevent the entrance of precipitation into the trench drains or flumes.</li>
</ol>
4. Plot Area Preparation
<ol>
	<li>Fill and tamp any minor voids on the upslope side of the wall using native topsoil from adjacent field areas.</li>
	<li>Use a small walk behind trencher to cut a 10 cm wide, 30 cm deep trench on the remaining 3 sides of all plots.
	<ol>
		<li>Insert 40 cm wide strips of 0.10 mm thick clear plastic vertically in the trenches to prevent lateral movement of water between plots.</li>
		<li>Install irrigation pipe and heads. Install six heads on 4.1 m2 spacing for each plot.</li>
		<li>Backfill and lightly tamp all trenches by hand. Mound the soil into a 5 cm tall by 30 cm wide berm over the trench area to prevent lateral movement of surface water between plots.</li>
		<li>Adjust irrigation heads to the top of the soil height in the berm areas.</li>
	</ol>
	</li>
	<li>Construct a diversion ditch to prevent upslope water from getting on the plots
	<ol>
		<li>Use a box blade to cut a V-shaped channel approximately 20 cm deep in the center and 2 m across. Note: The center of the channel should be approximately 1.25 m above the high side of the plot area and should extend across the upper side of all plots.</li>
		<li>Cut a sloped trench in the bottom of the channel. Note: To insure good drainage, the trench bottom should be 30 cm below the channel bottom at the high point in the center point above each block and have a minimum slope of 0.5% going to each end of each block. Trench bottoms must be hand smoothed and surveyed as necessary to ensure uniform slope.</li>
		<li>Add 5 cm of washed 6-9 mm pea gravel to the bottom of the trenches.</li>
		<li>Place a 15 cm diameter slotted drain line on the gravel surface and fill the trench with more 6-9 mm gravel.</li>
		<li>Cut trenches as needed at the ends and between the blocks of plots to route drainage water to discharge locations below the retaining wall. Use 15 cm diameter plain corrugated drain line and backfill these trenches with the excavated soil. Cover the trench and channel area with a layer of large 5-15 cm diameter bull rock.</li>
	</ol>
	</li>
</ol>
5. Planting and Initial Runoff Event
<ol>
	<li>Hand rake the plots to ensure a smooth seedbed with uniform slope in preparation for sod installation.</li>
	<li>Measure and document the slope of each plot using standard survey equipment by taking elevation measurements at distances of 0, 1.5, 3.0, 4.6, 6.1, and 7.6 m from the wall along the mid-line of each plot.</li>
	<li>Measure the depth of topsoil at 4 locations in each plot by inserting a 2.54 cm diameter soil probe into the soil until clay textured subsoil is encountered.</li>
	<li>Plant sod grown on a similar texture soil. Note: For this facility, mature &#8216;Raleigh&#8217; St. Augustinegrass (Stenotaphrum secundatum [Walt.] Kuntze) was used. However, other grasses may be used based on location, weather, and experimental design considerations. All plots may be sodded at one time or as in the present case, 12 plots (4 plots in each block) were planted on 08&#160;August 2012, with the remaining 12 plots planted on 12&#160;September 2012.</li>
	<li>Create a Runoff Event
	<ol>
		<li>Take initial readings of water meters and measure the soil moisture content of all plots.
		<ol>
			<li>Remove the lids from the valve boxes located at the head of each plot and record the initial water meter reading for each of the 24 plots.</li>
			<li>Using a portable hand held moisture probe, measure and record the soil moisture content of each plot. Note: For the initial characterization, 4 measurements were taken per plot (1 measurement in each quadrant of each plot) using 7.5 cm long probes. However, the number of measurements, length of probes, and type of instrument used may be varied based on the specific study objectives.</li>
		</ol>
		</li>
		<li>Program flow meters and samplers to measure flow and collect samples as desired. Note: 750 ml&#160;samples were collected after every 20 L of flow but other sample volumes and intervals may be used as appropriate.</li>
		<li>Operate the irrigation system for a predetermined time to apply sufficient water to cause runoff. Note: 20-21 mm of precipitation applied at a rate of 4.04 cm/hr was sufficient for this facility, however, this amount may vary based on site specific conditions.</li>
		<li>Record the ending water meter readings for each of the 24 plots. Collect irrigation water samples from the spray heads during operation. Label and transport runoff samples to the laboratory for analysis.</li>
	</ol>
	</li>
</ol>
6. Sample Analysis
<ol>
	<li>Measure the electrical conductivity and pH of the water samples by dipping probes directly into the samples. Then filter a 50 ml&#160;subsample of each water sample through a 0.7 &#181;m glass microfiber filter in preparation for chemical analysis.</li>
	<li>Measure dissolved organic carbon (DOC) and total dissolved nitrogen (TDN) using the USEPA method 415.115.
	<ol>
		<li>Make a 1,000 mg/L standard solution by adding 2.125 g dried potassium acid phthalate (1-KOCOC6H4-2-COOH) to a 1 L volumetric flask. Add approximately 500 ml&#160;distilled water, swirl to dissolve the chemical and bring to volume with distilled water. Store solution under refrigeration in a brown bottle.</li>
		<li>Make a 1,000 mg/L standard solution by adding 6.0677 g dried sodium nitrate to a 1 L volumetric flask. Add approximately 500 ml&#160;distilled water, swirl to dissolve the chemical, and bring to volume with distilled water.</li>
		<li>Make intermediate C and N standards that encompass the anticipated range of concentrations in the samples to be run by diluting subsamples of the standard solutions from steps 6.3.1-6.3.2.</li>
		<li>Pour approximately 16 ml&#160;of the water samples to be analyzed into a 24 ml&#160;sample vials and cover each with a septa and cap.</li>
		<li>Place filled vials in the autosampler tray keeping a record of what sample is in what position. Note: for quality assurance purposes a blank, two standards and two certified reference standards should be run after every 12th unknown.</li>
		<li>Place the autosampler tray in the machine and operate the auto analyzer following manufacturer&#8217;s instructions.</li>
	</ol>
	</li>
	<li>Measure phosphorus, nitrate and ammonia using the USEPA methods 365.1, 353.2, and 350.1, respectively, within 48 hr of sample collection16-18.
	<ol>
		<li>Make the following reagents and standards for phosphorus analysis:
		<ol>
			<li>Make a 5 N sulfuric acid stock solution by slowly adding 70 ml&#160;concentrated sulfuric acid to 400 ml&#160;distilled water in a 500 ml&#160;volumetric flask. Cool the solution to RT and dilute to volume using distilled water.</li>
			<li>Make a 0.3% potassium antimonyltartrate stock solution. Weigh 0.5 g antimony potassium tartrate, trihydrate C8H4K2O12Sb2&#8226;3H2O&#160;and dissolve it in about 50 ml&#160;distilled water in 100 ml&#160;volumetric flask. After it is dissolved, dilute to volume with distilled water and store at 4 &#176;C in a brown, glass-stoppered bottle.</li>
			<li>Make a 4% solution of ammonium molybdate by dissolving 4 g ammonium molybdate tetrahydrate, (NH4)6Mo7O24&#8226;4H2O,&#160;in 100 ml&#160;reagent water. Store in an acid washed plastic bottle at 4 &#176;C.</li>
			<li>Make a 15% w/w stock solution of sodium dodecyl sulfate (SDS). Dissolve 15 g of SDS CH3(CH2)11OSO3Na&#160;in 85 ml&#160;of distilled water. Note: This may require gentle stirring and heat to fully dissolve.</li>
			<li>Make a dilution SDS Solution (REAGENT 1) by adding 2 ml&#160;of 15% SDS stock solution to 98 ml&#160;distilled water. Cap flask and mix by inverting 5-6x.</li>
			<li>Make 100 ml&#160;of color reagent (REAGENT 2) by mixing the above reagents as follows: To 20 ml&#160;of distilled water add 50 ml&#160;of 5 N H2SO4 and mix. Add 5 ml&#160;of 0.3% antimony potassium tartrate solution and mix. Add 15 ml&#160;of 4% ammonium molybdate solution and mix. Add 10 ml&#160;of 15% w/w SDS solution and mix. Note: This solution can be stored in an acid washed bottle at RT for no more than one week.</li>
			<li>Make an ascorbic acid solution (REAGENT 3) by dissolving 0.88 g of ascorbic acid C6H8O6&#160;in 50 ml&#160;of distilled water. Add 0.5 ml&#160;of 15% SDS and swirl gently. Note: This solution must be prepared fresh daily.</li>
			<li>Make a 100 mg P/L standard solution by adding 0.4393 g dried KH2PO4&#160;to a 1 L volumetric flask. Add approximately 500 ml&#160;distilled water, swirl to dissolve the chemical and bring to volume with distilled water.</li>
		</ol>
		</li>
		<li>Make the following reagents and standards for nitrate analysis
		<ol>
			<li>Add 25 ml&#160;of concentrated phosphoric acid (H3PO4) to 150 ml&#160;distilled water in a 250 ml&#160;volumetric flask. Cool to RT and add 10.0 g sulfanilamide (4-NH2C6H4SO2NH2) and dissolve. Add 0.5 g N-(1-napthyl) ethylenediamine dihydrochloride (C10H7NHCH2CH2NH2&#8226;2HCl) and dissolve. Add 2 ml&#160;of concentrated rinse solution (from instrument manufacturer) and dilute to volume using distilled water. Note: Solution may be stored in a brown bottle for up to several weeks.</li>
			<li>Dissolve 85 g of ammonium chloride (NH4Cl) and 0.1 g disodium ethylenediamine tetraacetate (C10H14N2Na2O8&#8226;2H2O) in approximately 900 ml&#160;of distilled water in a 1 L volumetric flask. Adjust the pH to 8.5 by addition of concentrated ammonium hydroxide (NH4OH) and dilute to volume with distilled water.</li>
			<li>Put 200 ml&#160;of the solution from 6.4.2.2 in a 1 L volumetric and dilute to volume with distilled water. Adjust the pH to 8.5 by addition of concentrated ammonium hydroxide (NH4OH).</li>
			<li>Dissolve 7.218 g potassium nitrate (KNO3) in distilled water and dilute to 1&#160;L. Add 1 ml&#160;chloroform (CHCl3) as a preservative.</li>
		</ol>
		</li>
		<li>Make the following reagents and standards for ammonia analysis:
		<ol>
			<li>Dissolve 8 g sodium hydroxide (NaOH) in 125 ml&#160;of distilled water in a 250 ml&#160;volumetric flask. Cool to RT, add 20.75 g phenol (C6H5OH) and dissolve. Dilute to volume with distilled water and store up to 2 weeks in a brown bottle in the dark.</li>
			<li>Add 25 ml&#160;of bleach solution containing 5.25% NaOCl plus 0.5 ml&#160;of concentrated Probe Rinse Solution to a 50 ml&#160;volumetric flask. Dilute to volume with distilled water and mix.</li>
			<li>Dissolve 25 g EDTA disodium salt dihydrate (C10H14N2Na2O8&#8226;2H2O) and 2.75 g sodium hydroxide (NaOH) in approximately 450 ml&#160;distilled water in a 500 ml&#160;volumetric flask. Add 3 ml&#160;of concentrated Probe Rinse Solution, mix, and bring to volume with distilled water.</li>
			<li>Dissolve 0.075 g sodium nitroprusside dihydrate (Na2Fe(CN)5NO&#8226;2H2O) in 100 ml&#160;distilled water. Add 0.5 ml&#160;concentrated Probe Rinse Solution, mix and store in a brown bottle for up to 1 week.</li>
			<li>Make a 1,000 mg/L ammonia stock solution by dissolving 3.819 g dried anhydrous ammonium chloride (NH4Cl) in 500 ml&#160;distilled water and diluting to 1 L.</li>
		</ol>
		</li>
		<li>Place samples in 4 ml&#160;sample vials and cover each with a septa and cap.</li>
		<li>Place filled vials in the analyzer keeping a record of what sample is in what position. Note: for quality assurance purposes a certified reference standard should be run after every 12th unknown.</li>
		<li>Operate the analyzer following manufacturers instructions for the analyte of choice.</li>
	</ol>
	</li>
	<li>Measure cations (sodium, calcium, magnesium, and potassium) using Ion Chromatography.
	<ol>
		<li>Prepare a 1,000 mg/L stock solution of Na by adding 2.542 g NaCl to a 1 L volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 1,000 mg/L stock solution of K by adding 1.9070 g KCl to a 1 L volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 1,000 mg/L stock solution of Mg by adding 8.3608 g MgCl2&#8226;6H2O to a 1 L volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 1,000 mg/L stock solution of Ca by adding 3.6674 g CaCl&#8226;2H2O to a 1 L volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 350 mg/L working solution of Na by adding 35 ml&#160;of stock solution to a 100 ml&#160;volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 25 mg/L working solution of K by adding 2.5 ml&#160;of stock solution to a 100 ml&#160;volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 25 mg/L working solution of Mg by adding 2.5 ml&#160;of stock solution to a 100 ml&#160;volumetric flask and bring to volume with distilled water.</li>
		<li>Prepare a 75 mg/L working solution of Ca by adding 7.5 ml&#160;of stock solution to a 100 ml&#160;volumetric flask and bring to volume with distilled water.</li>
		<li>Refilter runoff water samples through a 0.2 &#956;m glass microfiber filter.</li>
		<li>Fill sample vial to fill line with sample or standard and seal with septa and cap.</li>
		<li>Place sample vials in the analyzer keeping track of sample locations. Note: for quality assurance purposes a blank&#160;and certified reference standard should be run after every 12th unknown.</li>
		<li>Operate the auto analyzer following manufacturer&#8217;s instructions.</li>
	</ol>
	</li>
</ol>

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W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+phosphorus+by+semi+automated+colorimetry.+Environmental+monitoring+systems+laboratory,+Office+of+research+and+development.">Determination of phosphorus by semi automated colorimetry. Environmental monitoring systems laboratory, Office of research and development.</a> U.S. Environmental Protection Agency. , (1993).</li><li>O'Dell, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+nitrate+nitrogen+by+semi+automated+colorimetry.+Revision+2.0+Edited+by+JW+O'Dell,+Environmental+monitoring+systems+laboratory.">Determination of nitrate nitrogen by semi automated colorimetry. Revision 2.0 Edited by JW O'Dell, Environmental monitoring systems laboratory.</a> Office of research and development, U.S. Environmental Protection Agency. , (1993).</li><li>O'Dell, J. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+ammonia+nitrogen+by+semi+automated+colorimetry.+Revision+2.0+Edited+by+JW+O'Dell,+Environmental+monitoring+systems+laboratory.">Determination of ammonia nitrogen by semi automated colorimetry. Revision 2.0 Edited by JW O'Dell, Environmental monitoring systems laboratory.</a> Office of research and development, U.S. Environmental Protection Agency. , (1993).</li><li>Gobel, P., Dierkes, C., Coldewey, W. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Storm+water+runoff+concentration+matrix+for+urban+areas.">Storm water runoff concentration matrix for urban areas.</a> J. Contam. Hydrol. 91, 26-42 (2007).</li><li>Vietor, D. M., Provin, T. L., White, R. H., Munster, C. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Runoff+losses+of+phosphorus+and+nitrogen+imported+in+sod+or+composted+manure+for+turf+establishment.">Runoff losses of phosphorus and nitrogen imported in sod or composted manure for turf establishment.</a> J. Env. Qual. 33, 358-366 (2004).</li></ol>

Except for S. Kelly being an employee of The Scotts Miracle-Gro Company, the authors declare that they have no competing financial interests.

Water flow over, into, and through soils is greatly affected by the topography, vegetative cover, and the soil physical properties. Excessively compacted soils and soils with high clay contents will exhibit reduced infiltration rates and increased amounts of runoff. Therefore, when building a facility of this nature, every effort should be made to use native soils with uniform slopes and minimize compaction from all types of traffic on the experimental areas during construction. In addition, compaction from post construc...

Four of the most rapidly growing, highly populated metropolitan areas are located in the southern U.S. in subtropical climates1. In addition, the largest percent change in urbanized land between 1982 and 1997 occurred in southern USA1. With increased urban areas comes a concomitant demand for potable water,&#160;much of which is used for outdoor use during summer months2. With new construction, programmable in-ground irrigation systems are often installed. Unfortunately, these systems are often programmed to deliver irrigation to urban landscaping more frequently and/or in volumes that exceed evapotranspiration demands of the landscape2. This results in a significant volume of runoff from urban landscaping to receiving waters, which contributes to what has been termed urban stream syndrome3. Symptoms of the urban stream syndrome include increased frequency of overland flow and erosive flow, increased nitrogen (N), phosphorus (P), toxicants, and temperature in addition to changes in channel morphology, freshwater biology, and ecosystem processes3.
Losses of N and P from agricultural ecosystems have been extensively studied and found to be primarily dependent on four factors: nutrient source, application rate, application timing, and nutrient placement4. While fewer published data currently exist on off site movement of nutrients from urban landscapes, these principals can be directly applied to turfgrass culture, whether in home lawns, sod farms, parks, or other green spaces. Additionally, improper irrigation practices which result in runoff from the landscape can exacerbate these losses.
Nutrient losses can be further altered by irrigation water quality. Areas in the southwest US often utilize more saline or sodic water for irrigation of home lawns and urban landscapes5,6. The chemical composition of the irrigation water may significantly alter soil chemistry causing a release of carbon, nitrogen, calcium, and other cations to runoff water. Recent work showed that increased sodium absorption ratio (SAR) of the extracting water significantly increased the amounts of carbon (C) and nitrogen (N) leached from St. Augustinegrass clippings, ryegrass clippings, and other organic materials7. Furthermore, water extractable soil C, N, and P losses from recreational turfgrass soils were significantly correlated with irrigation water chemical constituents6.
Washbusch et al. studied urban runoff in Madison, WI and found that lawns were the largest contributors of total phosphorus8. In addition, they also found that 25% of the total P in &#8220;Street Dirt&#8221; originated from leaves and grass clippings. In a typical rural setting, leaf litter falls onto the ground and then decomposes slowly releasing nutrients back to the soil environment. However, in urban environments, significant quantities of nutrient-rich leaves and grass clippings may fall on or get washed or blown onto hardscapes such as driveways, sidewalks, and roadways, subsequently making their way into the streets where they contribute to &#8220;street dirt&#8221;, much of which gets washed directly into receiving waterways.
Urban landscape soils are often disturbed and highly compacted during construction, which can also increase amounts of runoff due to reduced infiltration rates9. Kelling and Peterson reported that both total runoff volume and the nutrient concentrations in runoff from home lawns are increased from lawns that are compacted or have severely disturbed soil profiles due to previous construction activities10. Edmondson et al. on the other hand, found that urban soils were less compacted compared to surrounding agricultural soils in the urban and suburban region of Leicester, UK11. They attributed this to heavy agricultural machinery used, but they also noted that lawns had a greater soil bulk density than soil under trees and shrubs which was attributed to grass mowing and greater human trampling.
It would appear that in many situations, urban and suburban stream syndromes are significantly impacted by runoff and point-source discharges3,12. While point-sources can be manipulated through permits and recycling, additional research is needed to develop and test best management procedures for home lawn establishment and management to minimize nutrient losses to runoff. Past research efforts in this regard have often been centered along coastal areas where there are high sand content soils, due to concerns related to the effects of leaching and runoff losses of nutrients to coastal waters. However, when working with very sandy soils, one must have steep slopes and high rainfall rates to be able to generate any runoff13,14. In contrast, many of the soils in the central United States are fine textured and have low infiltration rates that result in significant amounts of runoff from even small rainfall events. Thus, it was desired to design and construct a runoff facility on native soil and slope typical of those that may occur on residential landscapes.
This paper describes the design, construction and function of a 1,000 m2 facility containing 24 individual 33.6 m2 field plots for measuring total runoff volumes at relatively small temporal resolutions and simultaneous collection of runoff water subsamples at selected volumetric or temporal intervals for measurement and quantification of chemical constituents of the runoff water.

<table border="0" cellpadding="0" cellspacing="0" width="778">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Flow meter</td>
			<td style="width:151px;">Teledyne Isco</td>
			<td style="width:216px;">Model 4230</td>
			<td style="width:167px;">Bubbling flow meter that measures and records water flow through flume</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Portable Sampler</td>
			<td>Teledyne Isco</td>
			<td>Model 6712</td>
			<td>Works in conjunction with the flow meter to collect water samples at predetermined intervals.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Flow Link Software to collect data</td>
			<td>Teledyne Isco</td>
			<td>Ver 5.0</td>
			<td>Allows communication between flow meter and computer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Pre-sloped trench drain</td>
			<td>Zurn Industries, LLC</td>
			<td>Z-886</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Irrigation Controller</td>
			<td>Toro Company</td>
			<td>VP Satellite</td>
			<td>Controls irrigation to each plot individually</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Electric Valves</td>
			<td>Hunter</td>
			<td>2.5 cm PGV</td>
			<td>Opens or closes water flow to individual plots based on signal from irrigation controller</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Spray nozzles</td>
			<td>RainBird</td>
			<td>HE-Van 12</td>
			<td>Sprays irrigation water in predetermined pattern and rate</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Irrigation heads</td>
			<td>Hunter</td>
			<td>Pro Spray 4</td>
			<td>4 inch pop up spray heads</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">6 inch slotted drain pipe</td>
			<td>Advanced Drainage Systems</td>
			<td>6410100</td>
			<td>single wall corregated HDPE - slotted</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">6 inch plain drain pipe</td>
			<td>Advanced Drainage Systems</td>
			<td>6400100</td>
			<td>single wall corregated HDPE - plain</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Filter Paper</td>
			<td>Whatman GF/F</td>
			<td>1825-047</td>
			<td>47mm diameter, binder-free, glass microfiber filter</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">pH Meter</td>
			<td>Fisher</td>
			<td>Accumet XL20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Combination pH probe</td>
			<td>Fisher</td>
			<td>13-620-130</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Automatic Temperature Compensating Probe</td>
			<td>Fisher</td>
			<td>13-602-19</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Electrical conductivity probe</td>
			<td>Fisher</td>
			<td>13-620-100</td>
			<td>Cell constant of 1.0</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">&#160;TOC-VCSH with total nitrogen unit TMN-1</td>
			<td>Shimadzu Corp</td>
			<td>TOC-VCSH with TMN-1</td>
			<td>dissolved C and N analyzer</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Smartchem 200</td>
			<td>Unity Scientific</td>
			<td>200</td>
			<td>Discrete Analyzer for P measurement</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">ICS 1000</td>
			<td>Dionex</td>
			<td>ICS 1000</td>
			<td>Ion Chromatography for Ca, Mg, K and Na measurment</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Portable Soil Moisture Meter</td>
			<td>Spectrum&#160;</td>
			<td style="width:216px;">FieldScout TDR 300</td>
			<td>7.5 cm long probes</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Totallizing Water Meters</td>
			<td style="width:151px;">Badger</td>
			<td style="width:216px;">3/4 inch water meters</td>
			<td>standard homeowner water meters</td>
		</tr>
	</tbody>
</table>

design and construction of an urban runoff research facility

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use due to increased demand for landscape irrigation. Improper irrigation practices and large rainfall events can result in runoff from urban landscapes which has potential to carry nutrients and sediments into local streams and lakes where they may contribute to eutrophication. A 1,000 m2 facility was constructed which consists of 24 individual 33.6 m2 field plots, each equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated urban landscapes. Runoff volumes from the first and second trials had coefficient of variability (CV) values of 38.2 and 28.7%, respectively. CV values for runoff pH, EC, and Na concentration for both trials were all under 10%. Concentrations of DOC, TDN, DON, PO4-P, K+, Mg2+, and Ca2+ had CV values less than 50% in both trials. Overall, the results of testing performed after sod installation at the facility indicated good uniformity between plots for runoff volumes and chemical constituents. The large plot size is sufficient to include much of the natural variability and therefore provides better simulation of urban landscape ecosystems.

Plot characteristics 
The average slope for all 24 plots was 3.7% and ranged from a low of 3.2% for plot 17 to a high of 4.1% for plot 2 (Table 1). Average topsoil thickness was 36 cm and ranged from a low of 25.0 cm for plot 24 to a high of 51.5 cm for plot 10 (Table 1).
Runoff volumes 
Runoff volumes from the first trial on 09&#160;August 2012 had a mean of 213.5 L and ranged from a low of 95.6 L to a high of 391 L with a coefficient of variability...

Watch this Scientific Journal Video about Design and Construction of an Urban Runoff Research Facility at JoVE.com

Design and Construction of an Urban Runoff Research Facility

This paper describes the design, construction, and function of a 1,000 m2 facility containing 24 individual 33.6 m2 field plots equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated home lawns.

As the urban population increases, so does the area of irrigated urban landscape. Summer water use in urban areas can be 2-3x winter base line water use ...

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13.0K Views.  Texas A&M University. This paper describes the design, construction, and function of a 1,000 m2 facility containing 24 individual 33.6 m2 field plots equipped for measuring total runoff volumes with time and collection of runoff subsamples at selected intervals for quantification of chemical constituents in the runoff water from simulated home lawns.

Video: Design and Construction of an Urban Runoff Research Facility

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 ...

A Protocol for Conducting Rainfall Simulation to Study Soil Runoff

Rainfall is a driving force for the transport of environmental contaminants from agricultural soils to surficial water bodies via surface runoff. The objective of this study was to characterize the effects of antecedent soil moisture content on the fate and transport of surface applied commercial urea, a common form of nitrogen (N) fertilizer, following a rainfall event that occurs within 24&#160;hr&#160;after fertilizer application. Although urea is assumed to be readily hydrolyzed to ammonium and therefore not often available for transport, recent studies suggest that urea can be transported from agricultural soils to coastal waters where it is implicated in harmful algal blooms. A rainfall simulator was used to apply a consistent rate of uniform rainfall across packed soil boxes that had been prewetted to different soil moisture contents. By controlling rainfall and soil physical characteristics, the effects of antecedent soil moisture on urea loss were isolated. Wetter soils exhibited shorter time from rainfall initiation to runoff initiation, greater total volume of runoff, higher urea concentrations in runoff, and greater mass loadings of urea in runoff. These results also demonstrate the importance of controlling for antecedent soil moisture content in studies designed to isolate other variables, such as soil physical or chemical characteristics, slope, soil cover, management, or rainfall characteristics. Because rainfall simulators are designed to deliver raindrops of similar size and velocity as natural rainfall, studies conducted under a standardized protocol can yield valuable data that, in turn, can be used to develop models for predicting the fate and transport of pollutants in runoff.

A rainfall simulator was used to apply a consistent rate of uniform rainfall to packed soil boxes in a study of the fate and transport of urea, a nonpoint source environmental contaminant. Under uniform soil and rainfall conditions, antecedent soil moisture content exerted strong control over urea loss in surface runoff.

Rainfall is a driving force for the transport of environmental contaminants from agricultural soils to surficial water bodies via surface runoff. The ...

A CO2 Concentration Gradient Facility for Testing CO2 Enrichment and Soil Effects on Grassland Ecosystem Function

Continuing increases in atmospheric carbon dioxide concentrations (CA) mandate techniques for examining impacts on terrestrial ecosystems. Most experiments examine only two or a few levels of CA concentration and a single soil type, but if CA can be varied as a gradient from subambient to superambient concentrations on multiple soils, we can discern whether past ecosystem responses may continue linearly in the future and whether responses may vary across the landscape. The Lysimeter Carbon Dioxide Gradient Facility applies a 250 to 500 &#181;l L-1 CA gradient to Blackland prairie plant communities established on lysimeters containing clay, silty clay, and sandy soils. The gradient is created as photosynthesis by vegetation enclosed in in temperature-controlled chambers progressively depletes carbon dioxide from air flowing directionally through the chambers. Maintaining proper air flow rate, adequate photosynthetic capacity, and temperature control are critical to overcome the main limitations of the system, which are declining photosynthetic rates and increased water stress during summer. The facility is an economical alternative to other techniques of CA enrichment, successfully discerns the shape of ecosystem responses to subambient to superambient CA enrichment, and can be adapted to test for interactions of carbon dioxide with other greenhouse gases such as methane or ozone.

A CO2 Concentration Gradient Facility for Testing CO2 Enrichment and Soil Effects on Grassland Ecosystem Function

The Lysimeter Carbon Dioxide Gradient Facility creates a 250 to 500 &#181;l L-1 linear carbon dioxide gradient in temperature-controlled chambers housing grassland plant communities on clay, silty clay, and sandy soil monoliths. The facility is used to determine how past and future carbon dioxide levels affect grassland carbon cycling.

Continuing increases in atmospheric carbon dioxide concentrations (CA) mandate techniques for examining impacts on terrestrial ecosystems. Most ...

Spotting Cheetahs: Identifying Individuals by Their Footprints

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges widely over sub-Saharan Africa and in parts of the Middle East. Cheetah conservationists face two major challenges, conflict with landowners over the killing of domestic livestock, and concern over range contraction. Understanding of the latter remains particularly poor2. Namibia is believed to support the largest number of cheetahs of any range country, around 30%, but estimates range from 2,9053 to 13,5204. The disparity is likely a result of the different techniques used in monitoring.
Current techniques, including invasive tagging with VHF or satellite/GPS collars, can be costly and unreliable. The footprint identification technique5 is a new tool accessible to both field scientists and also citizens with smartphones, who could potentially augment data collection. The footprint identification technique analyzes digital images of footprints captured according to a standardized protocol. Images are optimized and measured in data visualization software. Measurements of distances, angles, and areas of the footprint images are analyzed using a robust cross-validated pairwise discriminant analysis based on a customized model. The final output is in the form of a Ward's cluster dendrogram. A user-friendly graphic user interface (GUI) allows the user immediate access and clear interpretation of classification results.
The footprint identification technique algorithms are species specific because each species has a unique anatomy. The technique runs in a data visualization software, using its own scripting language (jsl) that can be customized for the footprint anatomy of any species. An initial classification algorithm is built from a training database of footprints from that species, collected from individuals of known identity. An algorithm derived from a cheetah of known identity is then able to classify free-ranging cheetahs of unknown identity. The footprint identification technique predicts individual cheetah identity with an accuracy of &gt;90%.

The cheetah (Acinonyx jubatus) is an iconic, endangered species, but conservation efforts are challenged by habitat shrinkage and conflict with commercial farmers. The footprint identification technique, a robust, accurate and cost-effective image classification system, is a new approach to monitoring cheetahs.

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges ...

Vegetated Treatment Systems for Removing Contaminants Associated with Surface Water Toxicity in Agriculture and Urban Runoff

Urban stormwater and agriculture irrigation runoff contain a complex mixture of contaminants that are often toxic to adjacent receiving waters. Runoff may be treated with simple systems designed to promote sorption of contaminants to vegetation and soils and promote infiltration. Two example systems are described: a bioswale treatment system for urban stormwater treatment, and a vegetated drainage ditch for treating agriculture irrigation runoff. Both have similar attributes that reduce contaminant loading in runoff: vegetation that results in sorption of the contaminants to the soil and plant surfaces, and water infiltration. These systems may also include the integration of granulated activated carbon as a polishing step to remove residual contaminants. Implementation of these systems in agriculture and urban watersheds requires system monitoring to verify treatment efficacy. This includes chemical monitoring for specific contaminants responsible for toxicity. The current paper emphasizes monitoring of current use pesticides since these are responsible for surface water toxicity to aquatic invertebrates.

This article summarizes the design attributes and the effectiveness of treatment systems that treat urban stormwater and agriculture irrigation runoff to remove pesticides and other contaminants associated with aquatic toxicity.

Urban stormwater and agriculture irrigation runoff contain a complex mixture of contaminants that are often toxic to adjacent receiving waters. Runoff may ...

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are also a useful scientific model to study motion vision and flight dynamics in a controlled laboratory setting. As the larvae develop in organically polluted water, they are useful models for investigating investment in microbial immunity. While large scale commercial breeding for agriculture already occurs, there are no standardized protocols for maintaining captive populations for scientific studies. This is important as commercial captive breeding programs focusing on mass output during peak pollination periods may fail to provide a population that is consistent, stable and robust throughout the year, as is often needed for other research purposes. Therefore, a method to establish, maintain and refresh a captive research population is required. Here, we describe the utilization of an artificial hibernation cycle, in addition to the nutritional and housing requirements, for long term maintenance of Eristalis tenax. Using these methods, we have significantly increased the health and longevity of captive populations of E. tenax compared to previous reports. We furthermore discuss small scale rearing methods and options for optimizing yields and manipulating population demographics.

Rearing and Long-Term Maintenance of Eristalis tenax Hoverflies for Research Studies

The overall goal of these procedures is to establish, maintain and refresh a captive population of Eristalis tenax in a research setting.

With an estimated 6000 species worldwide, hoverflies are ecologically important as alternative pollinators to domesticated honeybees. However, they are ...

Design and Use of an Apparatus for Quantifying Bivalve Suspension Feeding at Sea

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves (i.e., mussels, oysters, and clams) presents new challenges. Quantitative data on the feeding behavior of suspension-feeding mollusks is necessary to determine important ecosystem interactions of offshore shellfish farms, including their carrying capacity, the competition with the zooplankton community, the availability of trophic resources at different depths, and the deposition to the benthos. The biodeposition method is used to quantify feeding variables in suspension-feeding bivalves in a natural setting and represents a more realistic proxy than laboratory experiments. This method, however, relies upon a stable platform to satisfy the requirements that water flow rates supplied to the shellfish remain constant and the bivalves are undisturbed. A flow-through device and process for using the biodeposition method to quantify the feeding of bivalve mollusks were modified from a land-based format for shipboard use by building a two-dimensional gimbal table around the device. Planimeter data reveal a minimal pitch and yaw of the chambers containing the test shellfish despite boat motion, the flow rates within the chambers remain constant, and operators are able to collect the biodeposits (feces and pseudofeces) with sufficient consistency to obtain accurate measurements of the bivalve clearance, filtration, selection, ingestion, rejection, and absorption at offshore shellfish aquaculture sites.

A flow-through device for using the biodeposition method to quantify filtration and feeding behavior of bivalve mollusks was modified for shipboard use. A two-dimensional gimbal table built around the device isolates the apparatus from boat motion, thereby allowing the accurate quantification of bivalve filtration variables at offshore shellfish aquaculture sites.

As shellfish aquaculture moves from coastal embayments and estuaries to offshore locations, the need to quantify ecosystem interactions of farmed bivalves ...

Construction of a Low-cost Mobile Incubator for Field and Laboratory Use

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water quality. However, commercially available incubators are often costly, difficult to transport, not flexible in terms of volume, and/or poorly adapted to local field conditions where access to electricity is unreliable. The purpose of this study was to develop an adaptable, low-cost and transportable incubator that can be constructed using readily available components. The electronic core of the incubator was first developed. These components were then tested under a range of ambient temperature conditions (3.5 &#176;C - 39 &#176;C) using three types of incubator shells (polystyrene foam box, hard cooler box, and cardboard box covered with a survival blanket). The electronic core showed comparable performance to a standard laboratory incubator in terms of the time required to reach the set temperature, inner temperature stability and spatial dispersion, power consumption, and microbial growth. The incubator set-ups were also effective at moderate and low ambient temperatures (between 3.5 &#176;C and 27 &#176;C), and at high temperatures (39 &#176;C) when the incubator set temperature was higher. This incubator prototype is low-cost (&lt; 300 USD) and adaptable to a variety of materials and volumes. Its demountable structure makes it easy to transport. It can be used in both established laboratories with grid power or in remote settings powered by solar energy or a car battery. It is particularly useful as an equipment option for field laboratories in areas with limited access to resources for water quality monitoring.

This paper describes a method for building an adaptable, low-cost and transportable incubator for microbial testing of drinking water. Our design is based on widely available materials and can operate under a range of field conditions, while still offering the advantages of higher-end laboratory-based models.

Incubators are essential for a range of culture-based microbial methods, such as membrane filtration followed by cultivation for assessing drinking water ...

Combining Eye-tracking Data with an Analysis of Video Content from Free-viewing a Video of a Walk in an Urban Park Environment

As individuals increasingly live in cities, methods to study their everyday movements and the data that can be collected becomes important and valuable. Eye-tracking informatics are known to connect to a range of feelings, health conditions, mental states and actions. But because vision is the result of constant eye-movements, teasing out what is important from what is noise is complex and data intensive. Furthermore, a significant challenge is controlling for what people look at compared to what is presented to them.
The following presents a methodology for combining and analyzing eye-tracking on a video of a natural and complex scene with a machine learning technique for analyzing the content of the video. In the protocol we focus on analyzing data from filmed videos, how a video can be best used to record participants&#39; eye-tracking data, and importantly how the content of the video can be analyzed and combined with the eye-tracking data. We present a brief summary of the results and a discussion of the potential of the method for further studies in complex environments.

The objective of the protocol is to detail how to collect video data for use in the laboratory; how to record eye-tracking data of participants looking at the data and how to efficiently analyze the content of the videos that they were looking at using a machine learning technique.

As individuals increasingly live in cities, methods to study their everyday movements and the data that can be collected becomes important and valuable. ...

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.