The authors acknowledge the staff at the NCDA Sandhills Research Station for assistance with lysimeter installation and exhumation. Funding for experiments described in Representative Results was provided by the Center for Turfgrass Environmental Research &#38; Education. Video and manuscript production was supported by the North Carolina State University Departments of Soil Science and Crop Science.

Field sampling is performed in this experiment and is under the authorization of the North Carolina Department of Agriculture &#38; Consumer Services.
1. Field Lysimeter Installation
<ol>
	<li>Choose an experimental site on which lateral movement of applied chemicals is unlikely (i.e. sites with little or no slope). Select sites based on soil and vegetation properties of interest.</li>
	<li>If plots are vegetated, pull vegetation plugs prior to lysimeter installation (Figure 1A).</li>
	<li>Drive the lysimeters downward into the desired plots (with or without vegetation) using an inverted post driver, leaving ~1-2 cm of the lysimeter above the soil surface to contain the applied chemical and minimize lateral chemical movement. For this, use rolled and welded eighteen-gauge steel sheets (91 cm depth x 15 cm diameter) (Figure 1B). Use lysimeters of different materials and dimensions to fit research objectives.</li>
	<li>Replace vegetation plugs following lysimeter installation.</li>
	<li>Manage any vegetation as appropriate for the experiment. If plots are to remain bare, use spot applications of glyphosate to keep the areas free of vegetation.</li>
	<li>Ensure that irrigation, fertilization, and any other management practices are identical in the bareground and vegetated plots. Predetermine irrigation to meet research objectives.</li>
</ol>
2. Porewater Sampler Installation
<ol>
	<li>Install porewater samplers, such as PTFE/quartz (50/50%), in the middle of lysimeters to collect percolating porewater.</li>
	<li>Place a 2.5 cm stainless steel rod in the center of the lysimeter and insert it into the ground with a mallet to the desired sampler depth. 
	NOTE: An auger may also be used for this step.</li>
	<li>Prepare a silica flour and water slurry with 700 ml of irrigation water to ~900 g of chemically inert silica flour. Mix the slurry thoroughly before each sampler is placed in the mixture. Apply pressure between -50 to -70 kPa to the sampler from a handheld or battery-powered vacuum pump.</li>
	<li>Remove the sampler from the silica flour slurry after 10 min, and thoroughly mix the silica slurry again. Pour 60 ml of the slurry through a funnel connected to a 2.5 cm diameter pipe into the bottom of the hole.</li>
	<li>Place the sampler in the hole at the desired sampling depth with a plastic or metal pipe. Ensure that tubing from the sampler extends out of the hole. Use a slurry of nontreated, native soil and water to backfill the remaining hole.</li>
	<li>Allow time during backfilling for soil to settle; use a pipe to tamp added soil as needed.</li>
	<li>Backfill soil to the original level. If appropriate, replace vegetation at the top of the hole.</li>
	<li>Attach sampler tubing to a vacuum bottle via a section of fluorinated ethylene propylene (FEP) tubing. With a plastic tube clamp, connect a second tubing line out of the vacuum bottle to a vacuum pump.</li>
	<li>Cover tubing and collection bottles with black plastic or tape if the chemical(s) of interest is prone to photodegradation (Figure 1C).</li>
	<li>Apply vacuum pressure of approximately -50 to -70 kPa via the vacuum bottle to the samplers repeatedly over the course of several days before experimentation to ensure proper sampler installation.</li>
</ol>
3. Chemical Application to Lysimeters
<ol>
	<li>Allow at least two weeks for acclimation before chemical applications are made.</li>
	<li>Collect background porewater samples before lysimeter treatment to quantify background concentrations of the chemical(s) of interest.</li>
	<li>Apply the chemical of interest to the soil or vegetation by typical methods, such as with a hand-held CO2-pressurized boom sprayer (Figure 1D) or by distributing the granular formulation directly on the surface of the plot containing the lysimeter. If multiple chemical applications are necessary for effectiveness, apply them per typical use patterns or label directions. Leave some lysimeters untreated to serve as a control.</li>
</ol>
4. Porewater Collection and Analysis
<ol>
	<li>Apply approximately -50 to -70 kPa of vacuum to the porewater sampler vacuum bottles the day before or the day of sampling. Water surrounding the sampler will be drawn up through the sampler into the tubing, flowing to the vacuum bottle where it is collected until sampled. The soil volume from which porewater is collected and the water collection time may depend on factors such as soil type, soil texture, soil moisture content, and sampler depth.</li>
	<li>Collect samples at specified time intervals following chemical application, as predetermined by the researcher.</li>
	<li>Measure the volume of water collected into a graduated cylinder for each porewater sampler. If filtration is necessary, place the water in a Luer-Lok syringe (size will depend on volume of water) and pass sample through a 25 mm 0.2 &#181;m nylon filter.</li>
	<li>If different sample preservation methods are required and sufficient sample is collected, divide the sample into unique containers.</li>
	<li>Use a handheld pH meter to determine the pH of non-acidified samples.</li>
	<li>Adjust the pH by adding an adequate volume of the appropriate acid if needed for sample preservation. 
	NOTE: Concentrated acids can be corrosive or oxidizers and care should be taken when using them.</li>
	<li>Place samples on ice in a cooler or put in a refrigerator until analysis. Use analytical methods for chemical measurement such as inductively coupled plasma-mass spectrometry (ICP-MS), inductively coupled plasma-optical emission spectroscopy (ICP-OES), atomic absorption spectroscopy (AAS), or high-performance liquid chromatography (HPLC) to analyze the samples.</li>
</ol>
5. Lysimeter Exhumation, Soil/Vegetation Collection and Analysis
<ol>
	<li>Exhume the lysimeters, containing soil and vegetation, at specified time intervals following chemical application. Exhume nontreated lysimeters at each sampling time to determine background chemical concentrations within the soil and vegetation.</li>
	<li>Exhume lysimeters utilizing barrel clamps attached to a tractor implement. Lower the bucket to a position which allows for the clamps to be placed onto the lysimeter&#8217;s exposed edge.</li>
	<li>Lift the implement causing the clamps to grasp the exposed edge, pulling the lysimeter column out of the soil (Figure 1E).</li>
	<li>Cap exhumed lysimeter ends with insulation sheets cut to the diameter of the lysimeters. Hold caps in place with gallon-size polyethylene bags inserted over the lysimeter ends, and secure bags with duct tape.</li>
	<li>Transport the lysimeters to a field laboratory for soil and vegetation sample division. Process nontreated lysimeters first to prevent contamination among lysimeters.</li>
	<li>Use a reciprocating saw equipped with a metal cutting blade to cut the lysimeter lengthwise on one side. Cut the columns from the bottom (zone of anticipated lower concentration) to top (zone of anticipated higher concentration) to ensure the soil at deeper depths is not contaminated by soil at shallower depths.</li>
	<li>Split open the lysimeter. Use metal dividing plates to separate discrete soil and vegetation sections. Choose soil depth increments based on the length of the lysimeter and research objectives.</li>
	<li>Use spoons or spatulas to excavate the sectioned soil and vegetation. Place each sample in an appropriately labeled polyethylene freezer bag. Do not collect soil directly in contact with the lysimeter.</li>
	<li>Follow the excavation protocol for each desired sample depth. Place sample bags in a cooler filled with ice and transport them to a lab. Store samples in a freezer until analysis.</li>
</ol>

<ol>
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A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Our Nutrient World. The challenge to produce more food &amp; energy with less pollution. Key Messages for Rio +20. , (2012).</li><li>Du, W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fate+and+Ecological+Effects+of+Decabromodiphenyl+Ether+in+a+Field+Lysimeter.">Fate and Ecological Effects of Decabromodiphenyl Ether in a Field Lysimeter.</a> Environmental Science and Technology. 47, 9167-9174 (2013).</li><li>Fuhr, F., Burauel, P., Mittelstaedt, W., Putz, T., Wanner, U. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Environmental fate and effects of pesticides. , 1-29 (2003).</li><li>Hire, D. 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(ed USGS) Tallahassee. , (1996).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Organic+arsenical+herbicides+(MSMA,+DSMA,+CAMA,+and+Cacodylic+Acid),+reregistration+eligibility+decision;+notice+of+availability.">Organic arsenical herbicides (MSMA, DSMA, CAMA, and Cacodylic Acid), reregistration eligibility decision; notice of availability.</a> Environmental Protection Agency, Federal Register Environmental Documents. , 1-70 (2006).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=EPA+(not+Araujo+as+stated+before)+Organic+Arsenicals;+Amendments+to+Terminate+Uses:+Amendment+to+Existing+Stocks+Provision.">EPA (not Araujo as stated before) Organic Arsenicals; Amendments to Terminate Uses: Amendment to Existing Stocks Provision.</a> Environmental Protection Agency) 18590-18591 Federal Registrar. 78, (2013).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Drinking+Water+Regulations;+Arsenic+and+Clarifications+to+Compliance+and+New+Source+Contaminants+Monitoring+Final+Rule.">Drinking Water Regulations; Arsenic and Clarifications to Compliance and New Source Contaminants Monitoring Final Rule.</a> Environmental Protection Agency. 66, (2001).</li><li>Winton, K., Weber, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+review+of+field+lysimeter+studies+to+describe+the+environmental+fate+of+pesticides.">A review of field lysimeter studies to describe the environmental fate of pesticides.</a> Weed Technology. 10, 202-209 (1996).</li><li>Bergstr&#246;m, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+lysimeters+to+estimate+leaching+of+pesticides+in+agricultural+soils.">Use of lysimeters to estimate leaching of pesticides in agricultural soils.</a> Environmental Pollution. 67, 325-347 (1990).</li><li>Byron, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lysimeters+promoted+for+pesticide+research.">Lysimeters promoted for pesticide research.</a> Environmental Science and Technology. 31, (1997).</li><li>. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Infographic:+Pesticide+Planet.">Infographic: Pesticide Planet.</a> Science. 341, 730-731 (2013).</li><li>Severson, R., Grigal, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soil+solution+concentrations:+effects+of+extraction+time+using+porous+ceramic+cups+under+constant+tension.">Soil solution concentrations: effects of extraction time using porous ceramic cups under constant tension.</a> Water Resources Bulletin. 12, 1161-1170 (1976).</li><li>Allaire, S. E., Roulier, S., Cessna, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantifying+preferential+flow+in+soils:+A+review+of+different+techniques.">Quantifying preferential flow in soils: A review of different techniques.</a> Journal of Hydrology. 378, 179-204 (2009).</li><li>Weiherm&#252;ller, L., Kasteel, R., Vanderborght, J., P&#252;z, T., Vereecken, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Soil+Water+Extraction+with+a+Suction+Cup.">Soil Water Extraction with a Suction Cup.</a> Valdose Zone Journal. 4, 899-907 (2005).</li><li>Jury, W. A., Fluhler, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Advances in Agronomy. 47, 141-201 (1992).</li></ol>

The authors have nothing to disclose.

Utilizing an integrated field lysimetry and porewater sampling approach allows researchers to assess spatial and temporal distributions of a wide variety of land-applied chemicals. The fate of chemicals in soils and vegetated systems may be controlled by a number of environmental processes and attributes, such as downward leaching, volatilization, hydrolysis, photolysis, microbial transformation/degradation, plant uptake, soil type, and soil pH16,17. Unlike greenhouse or laboratory-based experiments, results from the field-based approach described here are obtained with minimal disruption to the system of study and therefore may be extrapolated to other systems or settings18. Knowing the quantity of the chemical applied, the area of the lysimeter, the potential volatilization of the chemical, the amount measured in the dissolved and solid phases, and the bulk density of the soil allows for determination of chemical mass balance and loading limit estimations for the system of interest &#8211; valuable information for predicting potential environmental threats, such as chemical leaching to groundwater.
The protocol described here illustrates one way to conduct an experiment employing integrated field lysimetry and porewater sampling. Many parts of this method may be adapted by researchers to address their specific objectives. For example, lysimeter size and type should be considered when preparing an experiment, and choices should reflect the chemical, soil, and plant properties of interest17. Placement of lysimeters must also be considered to minimize variability in environmental conditions and slope across the experimental area. Management practices (mowing, fertilizing, harvesting, etc.) determine not only the size of the lysimeter, but may affect installation depths and practicality, and should be considered to mimic real-world practices17,19.
Many types of porewater samplers are commercially available, and they represent a relatively inexpensive way to collect soil water from different depths. The size of the sampler, depth, samplers per lysimeter, and frequency of sampling should be considered when designing experiments. If the porewater sampler chosen is not large enough, suction applied may only collect from the immediate vicinity and not cover the entire lysimeter area20. A suggested solution is to use porewater plates that would cover a larger surface area21, although this may require extensive and undesirable soil excavation to accommodate sampler installation and may also limit water flow below the depth of the sampler. Another concern with porewater sampling is that, depending on soil type, sampler installation and vacuum application may cause porewater to preferentially flow toward the sampler or along lysimeter walls rather than naturally through the system, potentially altering chemical distributions17,22. Finally, to properly evaluate downward chemical leaching, adequate temporal porewater sampling is needed to ensure the chemical of interest does not leach past the sampler at times not captured by the sampling routine23.
One of the primary purposes of field lysimetry is to quantify the downward leaching potential of applied chemicals. However, this approach intentionally limits the impact of natural subsurface lateral flow on chemical transport. To overcome this limitation, scientists investigating chemical fate and behavior may use soil probes to collect soil cores, which has both advantages and disadvantages over field lysimetry. Once the area of interest is treated, a hand-held or tractor-mounted probe removes cores from plots that are smaller in size than typical lysimeters, requiring less area for experimentation and allowing for faster sampling. However, a consequence of using a probe is that it may push vegetation, soil, or roots downward, potentially contaminating deeper depths, compacting soil, and altering bulk densities. Soil-probe techniques also provide less protection against cross-plot contamination due to runoff and lateral subsurface flow.
One caveat of field lysimetry and porewater sampling is that 100% recovery of the applied chemical is rare17. There are unknowns when completing this type of research in the field compared with greenhouse or laboratory environments where more control is achieved with respect to weather, soil properties, and plant growth; consequently, results may vary across experimental trials3. Research utilizing both field and laboratory approaches may provide the most comprehensive examination of processes impacting the fate of chemicals in the environment. Nevertheless, field lysimetry and porewater sampling provide powerful, well-established techniques for assessing potential environmental concerns associated with chemicals. In the future, more studies will likely be performed using these techniques in order to better understand the fate of chemicals in the face of maintaining an adequate food supply, ensuring proper disposal of wastes, and upholding high standards of environmental protection.

Potentially toxic chemicals are routinely applied to land from sources such as pesticides, fertilizers, sewage/biosolids, industrial wastes, and municipal wastes1,2. The fate of these chemicals &#8211; which may include nutrients, trace elements, organics, and their associated metabolites &#8211; is often not well understood3. If the chemicals are not managed properly, they have the potential to threaten human and environmental health through their transfer to and buildup in plants, surface water, and groundwater. With a global population that may reach 10 billion people by 2050, there are growing demands on waste management and food production2, and land application of many chemicals has been increasing3,4. Accordingly, research is needed that quantifies the transformations, mobility, loading limits, and overall environmental risks from chemicals that require land disposal or that we depend upon to enhance crop health and yield.
A number of strategies have been employed to evaluate threats from chemicals applied in the environment. Laboratory-based, model-system studies have been conducted to provide information about fundamental mechanisms controlling the mobility of chemicals in soils. When analyzing chemical fate in a laboratory, complete manipulation of the &#8220;environment&#8221; and inputs may be achieved, but these rarely match real-world environmental conditions5,6. Thus, extrapolating lab results to field settings may lead to inaccurate predictions about chemical threats. In contrast, broad field measurements have been used to define chemical behavior in the environment. However, conclusions about environmental fate from these measurements are often complicated due to the frequently low use rates (e.g. a few g A-1) of applied chemicals, as well as the complex interactions between hydrological and biogeochemical processes in the environment that regulate chemical distributions.
Lysimetry, including field lysimetry, has historically been used by soil and crop scientists to systematically evaluate the downward mobility of chemicals applied to soils and established vegetation. A lysimeter is a device made of metal or plastic that is placed into a soil of interest and is used to determine the fate of chemicals applied in known amounts to a confined area. Soil and vegetation samples collected from lysimeters can be used to assess the evolution of chemical distributions over time. Because field lysimetry is carried out under natural environmental conditions, results may be used to predict real-case scenarios derived from chemical applications to soil systems. Early lysimeter studies measured transpiration, moisture flow, and/or nutrient movement. Modern-day lysimeter studies measure pesticide and nutrient dissipation, pesticide movement, volatility, and mass balance, along with the aforementioned measurements3.
A limitation of traditional field lysimetry is that chemical mobility within a soil profile is largely defined by solid-phase measurements, while less attention is paid to dissolved chemical concentrations in water percolating through the soils &#8211; a critical component that may impact the potential for groundwater contamination from land-applied chemicals. Although leachate from the bottom of the lysimeters is sometimes collected for analysis, this approach limits depth resolution of porewater concentrations and typically requires significant soil excavation prior to experimentation. Instead, to obtain data about chemical concentrations in soil water, porewater samplers may be utilized in field settings. Porewater samplers are installed in soils to collect water from discrete, desired depths and only minimally disturb the soil system. Porewater samplers have been referred to by many names including lysimeters, suction cup lysimeters, or soil solution samplers, convoluting their distinction with the traditional field lysimeters described above. In this paper, we will use the term &#8220;porewater sampler&#8221; to alleviate confusion.
Here, we demonstrate an experimental approach that combines field lysimetry and porewater sampling for evaluating the downward leaching potential of chemicals applied to vegetated soil or bareground systems. Lysimetry has been a powerful tool used since the 1700s7, while ceramic porewater sampling has been used since the early 1960s8. Integration of these robust techniques allows for field determination of both solid- and dissolved-phase chemical concentration distributions while minimizing soil disturbance. This paper describes factors to consider when designing an experiment, including site selection, device installation, and sample collection. The approach is illustrated with an experiment that evaluated the fate of an organic arsenical pesticide applied to a bareground and an established turfgrass system. The techniques described can be adjusted as necessary to examine the fate of a wide variety of chemicals, thereby providing invaluable tools to researchers and policy-makers who seek to understand the environmental fate and behavior of land-applied chemicals.

<table border="0" cellpadding="0" cellspacing="0" width="1187">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Name of Material/ Equipment</td>
			<td style="width:168px;">Company</td>
			<td style="width:119px;">Catalog Number</td>
			<td style="width:656px;">Comments/Description</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:244px;">Prenart Super Quartz Samplers (PFTE/Quartz)</td>
			<td style="width:168px;">Prenart Equipment ApS</td>
			<td style="width:119px;">N/A</td>
			<td>Any samplers for&#160; trace metal analysis can be used (e.g. SoilMoisture Equipment Corp.)</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Prenart Installation Kit</td>
			<td style="width:168px;">Prenart Equipment ApS</td>
			<td style="width:119px;">N/A</td>
			<td>Contains all items necessary to install porewater samplers</td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:244px;">2 L collecting bottles</td>
			<td style="width:168px;">Prenart Equipment ApS</td>
			<td></td>
			<td style="width:656px;">Bottles can also be purchased from Fisher Scientific (02-923-2) or other laboratory supply companies, but fittings will need to be adjusted. Bottles can be covered with dark material if light sensitive</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Portable vacuum pump</td>
			<td style="width:168px;">Prenart Equipment ApS</td>
			<td style="width:119px;">N/A</td>
			<td>Vacuporter from Decagon Devices or other field battery-operated or hand vacuum pump may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">1 oz HDPE Nalgene Bottles</td>
			<td style="width:168px;">Fisher Scientific</td>
			<td style="width:119px;">03-313-4A</td>
			<td>Sample bottle type will depend on analyte of interest and may be glass</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Concentrated nitric acid</td>
			<td style="width:168px;">Fisher Scientific</td>
			<td style="width:119px;">A509-P212</td>
			<td>Oxidizing and corrosive-other acids may be needed for preservation and should be used with caution</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">25 mm 0.2 &#181;m nylon syringe filters</td>
			<td style="width:168px;">VWR</td>
			<td style="width:119px;">28145-487</td>
			<td>Other filter types and pore sizes may be used, dependent on the analyte of interest and analytical instrumentation</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">60 mL Luer-Lok syringes</td>
			<td style="width:168px;">Fisher Scientific</td>
			<td style="width:119px;">13-689-8</td>
			<td>Other sizes may be used depending on sample volume collected</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Portable pH meter</td>
			<td style="width:168px;">VWR</td>
			<td style="width:119px;">248481-A01</td>
			<td>Other pH meters can be used following calibration</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Graduated Cylinder</td>
			<td style="width:168px;">any</td>
			<td style="width:119px;">N/A</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:244px;">Field lysimeters (metal, plastic, etc.)</td>
			<td style="width:168px;">N/A</td>
			<td style="width:119px;">N/A</td>
			<td>Often these are constructed based on the researchers specifications</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Inverted Post Driver Tractor</td>
			<td style="width:168px;">N/A</td>
			<td style="width:119px;">N/A</td>
			<td>Any tractor can be used to install the lysimeters</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Handheld Boom Sprayer</td>
			<td style="width:168px;">N/A</td>
			<td style="width:119px;">N/A</td>
			<td>To apply the rate needed for application&#160;</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Polyethylene bags</td>
			<td style="width:168px;">Johnson &#38; Johnson</td>
			<td style="width:119px;">N/A</td>
			<td>Other brands may be used for soil storage</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:244px;">Reciprocating saw</td>
			<td style="width:168px;">Black &#38; Decker&#160;</td>
			<td style="width:119px;">N/A</td>
			<td>Any reciprocating saw can be used with a metal cutting attachment</td>
		</tr>
	</tbody>
</table>

integrated field lysimetry and porewater sampling for evaluation of chemical mobility in soils and established vegetation

Potentially toxic chemicals are routinely applied to land to meet growing demands on waste management and food production, but the fate of these chemicals is often not well understood. Here we demonstrate an integrated field lysimetry and porewater sampling method for evaluating the mobility of chemicals applied to soils and established vegetation. Lysimeters, open columns made of metal or plastic, are driven into bareground or vegetated soils. Porewater samplers, which are commercially available and use vacuum to collect percolating soil water, are installed at predetermined depths within the lysimeters. At prearranged times following chemical application to experimental plots, porewater is collected, and lysimeters, containing soil and vegetation, are exhumed. By analyzing chemical concentrations in the lysimeter soil, vegetation, and porewater, downward leaching rates, soil retention capacities, and plant uptake for the chemical of interest may be quantified. Because field lysimetry and porewater sampling are conducted under natural environmental conditions and with minimal soil disturbance, derived results project real-case scenarios and provide valuable information for chemical management. As chemicals are increasingly applied to land worldwide, the described techniques may be utilized to determine whether applied chemicals pose adverse effects to human health or the environment.

This method allows for the accumulation of data on the fate of chemicals applied to bareground and vegetated soil systems5,10. This approach was used to evaluate arsenic (As) downward leaching, absorption, and translocation into plants for bermudagrass (Cynodon dactylon) systems following application of the organic arsenical herbicide monosodium methyl arsenate (MSMA)9. Since the 1960s, MSMA has been used in non-cropland, turfgrass, and cotton production, but there is growing concern that applied As may leach downward through soils and contaminate groundwater11,12. The U.S. Environmental Protection Agency (EPA) is currently considering phasing out MSMA, pending additional scientific review13,14.
Following MSMA application to bareground and bermudagrass lysimeters, the majority of As was retained within soil solid phases and vegetation throughout 1-year experiments (Figure 2, Table 1). Within the soils, the highest solid-phase As concentrations were found at 0-2 cm depth. Arsenic concentrations in MSMA-treated lysimeter samples were elevated above nontreated samples to the 8-15 cm depth increment, and at deeper depths, differences in solid-phase As concentrations between treated and nontreated lysimeters were statistically insignificant using a 2-tailed t-test with unequal variance (p&#8805;0.05). Arsenic was also taken up into the vegetation, and although they varied over time, As concentrations in bermudagrass foliage from treated plots were always significantly higher than those from nontreated plots. Overall, up to 101% of the applied As was recovered in soil and vegetation solid phases from the bermudagrass-covered lysimeters, whereas a maximum of 66% of the As was recovered in the bareground lysimeter samples (Table 1).
Porewater As concentrations in MSMA-treated plots were dependent on depth within the soil profile (Figure 3). At 30 cm depth, dissolved-phase As concentrations exceeded the EPA&#8217;s 10 &#181;g/L maximum drinking-water contaminant limit15, with concentrations immediately increasing following MSMA application and then subsequently decreasing over time. In contrast, porewater collected from 76.2 cm depth in the soil profile had As concentrations that were similar to background levels and consistently below the EPA limit, indicating that applied As did not migrate below the bounds of the experimental system.
The study discussed here highlights many of the aforementioned lysimetry and porewater sampling experimental design considerations. The field area contained roughly no slope, and ~1.5 cm of the lysimeter was left above ground to help prevent cross-plot contamination issues, while also allowing for proper bermudagrass management. The field area was chosen due to its low organic matter and high sand content (88% sand, 7% silt, 5% clay), representing a &#8220;worst-case&#8221; leaching scenario with respect to soil texture and As retention potential9. Porewater samplers were selected so that they would fit within the lysimeters, and several weeks were allowed for system equilibration prior to chemical application. Finally, episodic porewater sampling was heavily focused at the early stages of experimentation, when downward leaching of applied chemicals was considered most probable.
<img alt="Figure 1" fo:content-width="5in" src="/files/ftp_upload/51862/51862fig1highres.jpg" width="500" /> 
Figure 1. Photographs depicting select steps in the installation of lysimeters and porewater samplers. (A) Vegetation plugs are removed prior to lysimeter installation. (B) Lysimeters are driven into the ground using an inverted post driver. (C) Covered 2-L vacuum bottles are used to collect water from porewater samplers. (D) Chemical of interest is applied to randomized lysimeter plots. (E) Lysimeter soil cores are exhumed with a tractor implement.
<img alt="Figure 2" fo:content-width="5in" src="/files/ftp_upload/51862/51862fig2highres.jpg" width="500" /> 
Figure 2. Depth profiles of total As concentrations in lysimeter soil and bermudagrass vegetation over time following MSMA application. Symbol depths represent soil and vegetation samples from the following depth increments: 0 = above-ground foliage; &#8212;1 = 0 to 2 cm depth; &#8212;3 = 2 to 4 cm; &#8212;6 = 4 to 8 cm depth; &#8212;11.5 = 8 to 15 cm depth; &#8212;22.5 = 15 to 30 cm depth; and &#8212;37.5 = 30 to 45 cm depth. Error bars denote the standard deviation of measurements from replicate samples and lysimeters. Asterisks represent samples for which the measured As concentrations in MSMA-treated lysimeters were significantly higher than concentrations from respective nontreated lysimeter samples. Figure modified from Matteson et al., 20149; see reference for additional details.
<img alt="Figure 3" fo:content-width="5in" src="/files/ftp_upload/51862/51862fig3highres.jpg" width="500" /> 
Figure 3. Porewater As concentrations from two depths (30 and 76.2 cm) within MSMA-treated, bermudagrass-covered lysimeters.
<table border="0" cellpadding="0" cellspacing="0">
	<tbody>
		<tr>
			<td>Days After MSMA Treatment</td>
			<td>Vegetated or Bare</td>
			<td>As Recovered in Soil (%)</td>
			<td>As Recovered in Vegetation (%)</td>
			<td>Total As Recovered (%)</td>
		</tr>
		<tr>
			<td>36</td>
			<td>Vegetated</td>
			<td>83</td>
			<td>10</td>
			<td>93</td>
		</tr>
		<tr>
			<td>36</td>
			<td>Bare</td>
			<td>62</td>
			<td>-</td>
			<td>62</td>
		</tr>
		<tr>
			<td>64</td>
			<td>Vegetated</td>
			<td>47</td>
			<td>3</td>
			<td>50</td>
		</tr>
		<tr>
			<td>64</td>
			<td>Bare</td>
			<td>60</td>
			<td>-</td>
			<td>60</td>
		</tr>
		<tr>
			<td>119</td>
			<td>Vegetated</td>
			<td>83</td>
			<td>9</td>
			<td>92</td>
		</tr>
		<tr>
			<td>119</td>
			<td>Bare</td>
			<td>66</td>
			<td>-</td>
			<td>66</td>
		</tr>
		<tr>
			<td>364</td>
			<td>Vegetated</td>
			<td>98</td>
			<td>4</td>
			<td>101</td>
		</tr>
		<tr>
			<td>364</td>
			<td>Bare</td>
			<td>55</td>
			<td>-</td>
			<td>55</td>
		</tr>
	</tbody>
</table>
Table 1. Calculated total As recoveries in lysimeter soil and bermudagrass vegetation following MSMA application. Recovery values represent total As in lysimeter samples from MSMA-treated plots minus total As in nontreated samples, all divided by the quantity of As added to the system via MSMA application. Table modified from Matteson et al., 20149; see reference for additional details.

Watch this Scientific Journal Video about Integrated Field Lysimetry and Porewater Sampling for Evaluation of Chemical Mobility in Soils and Established Vegetation at JoVE.com

Integrated Field Lysimetry and Porewater Sampling for Evaluation of Chemical Mobility in Soils and Established Vegetation

Field lysimetry and porewater sampling allow researchers to evaluate the fate of chemicals applied to soils and established vegetation. The goal of this protocol is to demonstrate how to install required instrumentation and collect samples for chemical analysis during integrated field lysimetry and porewater sampling experiments.

Potentially toxic chemicals are routinely applied to land to meet growing demands on waste management and food production, but the fate of these chemicals ...

integrated-field-lysimetry-porewater-sampling-for-evaluation-chemical

Nanyang Technological University

Research

JoVE Journal

Environment

14.2K Views.  North Carolina State University. Field lysimetry and porewater sampling allow researchers to evaluate the fate of chemicals applied to soils and established vegetation. The goal of this protocol is to demonstrate how to install required instrumentation and collect samples for chemical analysis during integrated field lysimetry and porewater sampling experiments.

Video: Integrated Field Lysimetry and Porewater Sampling for Evaluation of Chemical Mobility in Soils and Established Vegetation

Evaluation of Integrated Anaerobic Digestion and Hydrothermal Carbonization for Bioenergy Production

Lignocellulosic biomass is one of the most abundant yet underutilized renewable energy resources. Both anaerobic digestion (AD) and hydrothermal carbonization (HTC) are promising technologies for bioenergy production from biomass in terms of biogas and HTC biochar, respectively. In this study, the combination of AD and HTC is proposed to increase overall bioenergy production. Wheat straw was anaerobically digested in a novel upflow anaerobic solid state reactor (UASS) in both mesophilic (37 &#176;C) and thermophilic (55 &#176;C) conditions. Wet digested from thermophilic AD was hydrothermally carbonized at 230 &#176;C for 6 hr for HTC biochar production. At thermophilic temperature, the UASS system yields an average of 165 LCH4/kgVS&#160;(VS: volatile solids) and 121 L CH4/kgVS&#160;at mesophilic AD over the continuous operation of 200 days. Meanwhile, 43.4 g of HTC biochar with 29.6 MJ/kgdry_biochar was obtained from HTC of 1 kg digestate (dry basis) from mesophilic AD. The combination of AD and HTC, in this particular set of experiment yield 13.2 MJ of energy per 1 kg of dry wheat straw, which is at least 20% higher than HTC alone and 60.2% higher than AD only.

A novel Upflow Anaerobic Solid State (UASS) reactor was used for biogas production from fibrous feedstock. Digestate from UASS reactor was hydrothermally carbonized into HTC biochar in a pressurized batch reactor. The integration of the two bioenergy concepts was applied in this study to increase overall bioenergy production.

Lignocellulosic biomass is one of the most abundant yet underutilized renewable energy resources. Both anaerobic digestion (AD) and hydrothermal ...

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 Bending Test for Determining the Atterberg Plastic Limit in Soils

The thread rolling test is the most commonly used method to determine the plastic limit (PL) in soils. It has been widely criticized, because a considerable subjective judgment from the operator that carries out the test is involved during its performance, which may affect the final result significantly. Different alternative methods have been put forward, but they cannot compete with the standard rolling test in speed, simplicity and cost.
In an earlier study by the authors, a simple method with a simple device to determine the PL was presented (the &#34;thread bending test&#34; or simply &#34;bending test&#34;); this method allowed the PL to be obtained with minimal operator interference. In the present paper a version of the original bending test is shown. The experimental basis is the same as the original bending test: soil threads which are 3 mm in diameter and 52 mm long are bent until they start to crack, so that both the bending produced and its related moisture content are determined. However, this new version enables the calculation of PL from an equation, so it is not necessary to plot any curve or straight line to obtain this parameter and, in fact, the PL can be achieved with only one experimental point (but two experimental points are recommended).
The PL results obtained with this new version are very similar to those obtained through the original bending test and the standard rolling test by a highly experienced operator. Only in particular cases of high plasticity cohesive soils, there is a greater difference in the result. Despite this, the bending test works very well for all types of soil, both cohesive and very low plasticity soils, where the latter are the most difficult to test via the standard thread rolling method.

The traditional standardized test for determining the plastic limit in soils is performed by hand, and the result varies depending on the operator. An alternative method based on bending measurements is presented in this study. This allows the plastic limit to be obtained with a clear and objective criterion.

The thread rolling test is the most commonly used method to determine the plastic limit (PL) in soils. It has been widely criticized, because a ...

Extraction and Analysis of Microbial Phospholipid Fatty Acids in Soils

Phospholipid fatty acids (PLFAs) are key components of microbial cell membranes. The analysis of PLFAs extracted from soils can provide information about the overall structure of terrestrial microbial communities. PLFA profiling has been extensively used in a range of ecosystems as a biological index of overall soil quality, and as a quantitative indicator of soil response to land management and other environmental stressors.
The standard method presented here outlines four key steps: 1. lipid extraction from soil samples with a single-phase chloroform mixture, 2. fractionation using solid phase extraction columns to isolate phospholipids from other extracted lipids, 3. methanolysis of phospholipids to produce fatty acid methyl esters (FAMEs), and 4. FAME analysis by capillary gas chromatography using a flame ionization detector (GC-FID). Two standards are used, including 1,2-dinonadecanoyl-sn-glycero-3-phosphocholine (PC(19:0/19:0)) to assess the overall recovery of the extraction method, and methyl decanoate (MeC10:0) as an internal standard (ISTD) for the GC analysis.

Phospholipid fatty acids provide information about the structure of soil microbial communities. We present methods for extraction from soil samples with a single-phase chloroform mixture, fractionation of extracted lipids using solid phase extraction columns, and methanolysis to produce fatty acid methyl esters, which are analyzed by capillary gas chromatography.

Phospholipid fatty acids (PLFAs) are key components of microbial cell membranes. The analysis of PLFAs extracted from soils can provide information about ...

Modification and Application of a Leaf Blower-vac for Field Sampling of Arthropods

Rice fields host a large diversity of arthropods, but investigating their population dynamics and interactions is challenging. Here we describe the modification and application of a leaf blower-vac for suction sampling of arthropod populations in rice. When used in combination with an enclosure, application of this sampling device provides absolute estimates of the populations of arthropods as numbers per standardized sampling area. The sampling efficiency depends critically on the sampling duration. In a mature rice crop, a two-minute sampling in an enclosure of 0.13 m2 yields more than 90% of the arthropod population. The device also allows sampling of arthropods dwelling on the water surface or the soil in rice paddies, but it is not suitable for sampling fast flying insects, such as predatory Odonata or larger hymenopterous parasitoids. The modified blower-vac is simple to construct, and cheaper and easier to handle than traditional suction sampling devices, such as D-vac. The low cost makes the modified blower-vac also accessible to researchers in developing countries.

Assessment of arthropod abundance in crops is critical for investigating population dynamics and species interactions. Here we describe the modification and application of a leaf blower-vac for suction sampling of arthropods in rice.

Rice fields host a large diversity of arthropods, but investigating their population dynamics and interactions is challenging. Here we describe the ...

Methods of Soil Resampling to Monitor Changes in the Chemical Concentrations of Forest Soils

Recent soils research has shown that important chemical soil characteristics can change in less than a decade, often the result of broad environmental changes. Repeated sampling to monitor these changes in forest soils is a relatively new practice that is not well documented in the literature and has only recently been broadly embraced by the scientific community. The objective of this protocol is therefore to synthesize the latest information on methods of soil resampling in a format that can be used to design and implement a soil monitoring program. Successful monitoring of forest soils requires that a study unit be defined within an area of forested land that can be characterized with replicate sampling locations. A resampling interval of 5 years is recommended, but if monitoring is done to evaluate a specific environmental driver, the rate of change expected in that driver should be taken into consideration. Here, we show that the sampling of the profile can be done by horizon where boundaries can be clearly identified and horizons are sufficiently thick to remove soil without contamination from horizons above or below. Otherwise, sampling can be done by depth interval. Archiving of sample for future reanalysis is a key step in avoiding analytical bias and providing the opportunity for additional analyses as new questions arise.

Repeated soil sampling has recently been shown to be an effective way to monitor forest soil change over years and decades. To support its use, a protocol is presented that synthesizes the latest information on soil resampling methods to aid in the design and implementation of successful soil monitoring programs.

Recent soils research has shown that important chemical soil characteristics can change in less than a decade, often the result of broad environmental ...

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 ...

A New Portable In Vitro Exposure Cassette for Aerosol Sampling

This protocol introduces a new in vitro exposure system, capable of being worn, including its characterization and performance. Air-liquid interface (ALI) in vitro exposure systems are often large and bulky, making transport to the field and operation at the source of emission or within the breathing zone difficult. Through miniaturization of these systems, the lab can be brought to the field, expediting processing time and providing a more appropriate exposure method that does not alter the aerosol prior to contacting the cells. The Portable In vitro Exposure Cassette (PIVEC) adapts a 37 mm filter cassette to allow for in vitro toxicity testing outside of a traditional laboratory setting. The PIVEC was characterized using three sizes of copper nanoparticles to determine deposition efficiency based on gravimetric and particle number concentration analysis. Initial cytotoxicity experiments were performed with exposed lung cells to determine the ability of the system to deposit particles while maintaining cell viability. The PIVEC provides a similar or increased deposition efficiency when comparing to available perpendicular flow in vitro exposure devices. Despite the lower sample throughput, the small size gives some advantages to the current in vitro ALI exposure systems. These include the ability to be worn for personal monitoring, mobility from the laboratory to the source of emission, and the option to set-up multiple systems for spatial resolution while maintaining a lower user cost. The PIVEC is a system capable of collecting aerosols in the field and within the breathing zone onto an air-interfaced, in vitro model.

A New Portable In Vitro Exposure Cassette for Aerosol Sampling

Here, we present a protocol to perform portable cellular aerosol exposures and measure cellular response. The method uses cells, grown at the air-liquid interface, mimicking in vivo physiology. Cellular response to copper nanoparticle aerosols was observed as oxidative stress through reactive oxygen species generation and cytotoxicity as lactate dehydrogenase release.

This protocol introduces a new in vitro exposure system, capable of being worn, including its characterization and performance. Air-liquid interface (ALI) ...

Combined Size and Density Fractionation of Soils for Investigations of Organo-Mineral Interactions

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF relies on sequential density separation and sedimentation steps to isolate (1) the free light fraction (uncomplexed organic matter), (2) the occluded light fraction (uncomplexed organic matter trapped in soil aggregates) and (3) a variable number of heavy fractions (soil minerals and their associated organic matter) differing in composition. Provided that the parameters of the CSDF (dispersion energy, density cut-offs, sedimentation time) are properly selected, the method yields heavy fractions of relatively homogeneous mineral composition. Each of these fractions is expected to have a different complexing ability towards organic matter, rendering this a useful method to isolate and study the nature of organo-mineral interactions. Combining density and particle size separation brings an improved resolution compared to simple size or density fractionation methods, allowing the separation of heavy components according to both mineralogy and size (related to surface area) criteria. As is the case for all physical fractionation methods, it may be considered as less disruptive or aggressive than chemically-based extraction methods. However, CSDF is a time-consuming method and furthermore, the quantity of material obtained in some fractions can be limiting for subsequent analysis. Following CSDF, the fractions may be analyzed for mineralogical composition, soil organic carbon concentration and organic matter chemistry. The method provides quantitative information about organic carbon distribution within a soil sample and brings light to the sorptive capacity of the different, naturally-occurring mineral phases, thus providing mechanistic information about the preferential nature of organo-mineral interactions in soils (i.e., which minerals, what type of organic matter).

Combined size and density fractionation (CSDF) is a method to physically separate soil into fractions differing in texture (particle size) and mineralogy (density). The purpose is to isolate fractions with different reactivities towards soil organic matter (SOM), in order to better understand organo-mineral interactions and SOM dynamics.

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF ...

High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling in soils, to pathogen competition, plant growth promotion, and cross-kingdom signaling. Furthermore, siderophores are also of commercial interest in bioleaching and bioweathering of metal-bearing minerals and ores. A rapid, cost effective, and robust means of quantitatively assessing siderophore production in complex samples is key to identifying important aspects of the ecological ramifications of siderophore activity, including, novel siderophore producing microbes. The method presented here was developed to assess siderophore activity of in-tact microbiome communities, in environmental samples, such as soil or plant tissues. The samples were homogenized and diluted in a modified M9 medium (without Fe), and enrichment cultures were incubated for 3 days. Siderophore production was assessed in samples at 24, 48, and 72 hours (h) using a novel 96-well microplate CAS (Chrome azurol sulphonate)-Fe agar assay, an adaptation of the traditionally tedious and time-consuming colorimetric method of assessing siderophore activity, performed on individual cultivated microbial isolates. We applied our method to 4 different genotypes/Lines of wheat (Triticum aestivum L.), including Lewjain, Madsen, and PI561725, and PI561727 commonly grown in the inland Pacific Northwest. Siderophore production was clearly impacted by the genotype of wheat, and in the specific types of plant tissues observed. We successfully used our method to rapidly screen for the influence of plant genotype on siderophore production, a key function in terrestrial and aquatic ecosystems. We produced many technical replicates, yielding very reliable statistical differences in soils and within plant tissues. Importantly, the results show the proposed method can be used to rapidly examine siderophore production in complex samples with a high degree of reliability, in a manner that allows communities to be preserved for later work to identify taxa and functional genes.

We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling ...