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>

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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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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 f...

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><tbody><tr><td>Prenart Super Quartz Samplers (PFTE/Quartz)</td><td>Prenart Equipment ApS</td><td>N/A</td><td>Any samplers for&amp;nbsp; trace metal analysis can be used (&lt;em&gt;e.g.&lt;/em&gt; SoilMoisture Equipment Corp.)</td></tr><tr><td>Prenart installation kit</td><td>Prenart Equipment ApS</td><td>N/A</td><td>Contains all items necessary to install porewater samplers</td></tr><tr><td>2 L collecting bottles</td><td>Prenart Equipment ApS</td><td></td><td>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><td>Portable vacuum pump</td><td>Prenart Equipment ApS</td><td>N/A</td><td>Vacuporter from Decagon Devices or other field battery-operated or hand vacuum pump may be used</td></tr><tr><td>1 oz HDPE Nalgene bottles</td><td>Fisher Scientific</td><td>03-313-4A</td><td>Sample bottle type will depend on analyte of interest and may be glass</td></tr><tr><td>Concentrated nitric acid</td><td>Fisher Scientific</td><td>A509-P212</td><td>Oxidizing and corrosive-other acids may be needed for preservation and should be used with caution</td></tr><tr><td>25 mm 0.2 &amp;micro;m nylon syringe filters</td><td>VWR</td><td>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><td>60 ml Luer-Lok syringes</td><td>Fisher Scientific</td><td>13-689-8</td><td>Other sizes may be used depending on sample volume collected</td></tr><tr><td>Portable pH meter</td><td>VWR</td><td>248481-A01</td><td>Other pH meters can be used following calibration</td></tr><tr><td>Graduated cylinder</td><td>any</td><td>N/A</td><td></td></tr><tr><td>Field lysimeters (metal, plastic, &lt;em&gt;etc.&lt;/em&gt;)</td><td>N/A</td><td>N/A</td><td>Often these are constructed based on the researchers specifications</td></tr><tr><td>Inverted post driver tractor</td><td>N/A</td><td>N/A</td><td>Any tractor can be used to install the lysimeters</td></tr><tr><td>Handheld boom sprayer</td><td>N/A</td><td>N/A</td><td>To apply the rate needed for application&amp;nbsp;</td></tr><tr><td>Polyethylene bags</td><td>Johnson &amp;amp; Johnson</td><td>N/A</td><td>Other brands may be used for soil storage</td></tr><tr><td>Reciprocating saw</td><td>Black &amp;amp; Decker&amp;nbsp;</td><td>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 a...

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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 ...

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14.2K Views.  North Carolina State University.  Veld lysimetry en poriënwater bemonstering kunnen de onderzoekers het lot van chemische stoffen die de bodem en bestaande vegetatie te evalueren. Het doel van dit protocol is om aan te tonen hoe de benodigde instrumentatie installeren en monsters voor chemische analyse te verzamelen tijdens geïntegreerde veld lysimetry en poriënwater bemonstering experimenten. 

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

A Protocol for Collecting and Constructing Soil Core Lysimeters

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

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

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

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled Earth-system processes. Such knowledge is imperative in improving predictions of hydro-biogeochemical cycles, especially under climate change scenarios. We present an experimental method, designed to capture sub-surface heterogeneity of an initially homogeneous soil system. This method is based on destructive sampling of a soil lysimeter designed to simulate a small-scale hillslope. A weighing lysimeter of one cubic meter capacity was divided into sections (voxels) and was excavated layer-by-layer, with sub samples being collected from each voxel. The excavation procedure was aimed at detecting the incipient heterogeneity of the system by focusing on the spatial assessment of hydrological, geochemical, and microbiological properties of the soil. Representative results of a few physicochemical variables tested show the development of heterogeneity. Additional work to test interactions between hydrological, geochemical, and microbiological signatures is planned to interpret the observed patterns. Our study also demonstrates the possibility of carrying out similar excavations in order to observe and quantify different aspects of soil-development under varying environmental conditions and scale.

This study presents an excavation method for investigating subsurface hydrological, geochemical, and microbiological heterogeneity of a soil lysimeter. The lysimeter simulates an artificial hillslope which was initially under homogeneous condition and had been subjected to approximately 5,000 mm of water over eight cycles of irrigation in an 18-month period.

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled ...

A Lipid Extraction and Analysis Method for Characterizing Soil Microbes in Experiments with Many Samples

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial communities, a relatively precise but effort-intensive technique to assay microbial community composition is phospholipid fatty acid (PLFA) analysis. PLFA was developed to analyze phospholipid biomarkers, which can be used as indicators of microbial biomass and the composition of broad functional groups of fungi and bacteria. It has commonly been used to compare soils under alternative plant communities, ecology, and management regimes. The PLFA method has been shown to be sensitive to detecting shifts in microbial community composition.
An alternative method, fatty acid methyl ester extraction and analysis (MIDI-FA) was developed for rapid extraction of total lipids, without separation of the phospholipid fraction, from pure cultures as a microbial identification technique. This method is rapid but is less suited for soil samples because it lacks an initial step separating soil particles and begins instead with a saponification reaction that likely produces artifacts from the background organic matter in the soil. 
This article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples. The method combines the accuracy achieved through PLFA profiling by extracting and concentrating soil lipids as a first step, and a reduction in effort by saponifying the organic material extracted and processing with the MIDI-FA method as a second step.

The article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples.

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial ...

Measuring and Mapping Patterns of Soil Erosion and Deposition Related to Soil Carbonate Concentrations Under Agricultural Management

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonate (CaCO3) profiles. The objective is to describe a simple conceptual model and detailed protocol for repeatable field and laboratory measurements of these quantities. Here, accurate elevation is measured using a ground-based differential global positioning system (GPS); other data acquisition methods could be applied to the same basic method. Soil samples are collected from prescribed depth intervals and analyzed in the lab using an efficient and precise modified pressure-calcimeter method for quantitative analysis of inorganic carbon concentration. Standard statistical methods are applied to point data, and representative results show significant correlations between changes in soil surface layer CaCO3 and changes in elevation consistent with the conceptual model; CaCO3 generally decreased in depositional areas and increased in erosional areas. Maps are derived from point measurements of elevation and soil CaCO3 to aid analyses. A map of erosional and depositional patterns at the study site, a rain-fed winter wheat field cropped in alternating wheat-fallow strips, shows the interacting effects of water and wind erosion affected by management and topography. Alternative sampling methods and depth intervals are discussed and recommended for future work relating soil erosion and deposition to soil CaCO3.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonates. Repeatable methods for field and laboratory measurements of these quantities and data analysis methods are described here.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes ...

Single-throughput Complementary High-resolution Analytical Techniques for Characterizing Complex Natural Organic Matter Mixtures

Natural organic matter (NOM) is composed of a highly complex mixture of thousands of organic compounds which, historically, proved difficult to characterize. However, to understand the thermodynamic and kinetic controls on greenhouse gas (carbon dioxide [CO2] and methane [CH4]) production resulting from the decomposition of NOM, a molecular-level characterization coupled with microbial proteome analyses is necessary. Further, climate and environmental changes are expected to perturb natural ecosystems, potentially upsetting complex interactions that influence both the supply of organic matter substrates and the microorganisms performing the transformations. A detailed molecular characterization of the organic matter, microbial proteomics, and the pathways and transformations by which organic matter is decomposed will be necessary to predict the direction and magnitude of the effects of environmental changes. This article describes a methodological throughput for comprehensive metabolite characterization in a single sample by direct injection Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS), gas chromatography mass spectrometry (GC-MS), nuclear magnetic resonance (NMR) spectroscopy, liquid chromatography mass spectrometry (LC-MS), and proteomics analysis. This approach results in a fully-paired dataset which improves statistical confidence for inferring pathways of organic matter decomposition, the resulting CO2 and CH4 production rates, and their responses to environmental perturbation. Herein we present results of applying this method to NOM samples collected from peatlands; however, the protocol is applicable to any NOM sample (e.g., peat, forested soils, marine sediments, etc.).

This protocol describes a single throughput for complementary analytical and omics techniques culminating in a fully-paired characterization of natural organic matter and microbial proteomics in different ecosystems. This approach permits robust comparisons for identifying metabolic pathways and transformations important for describing greenhouse gas production and predicting responses to environmental change.

Natural organic matter (NOM) is composed of a highly complex mixture of thousands of organic compounds which, historically, proved difficult to ...

Measuring Phosphorus Release in Laboratory Microcosms for Water Quality Assessment

Phosphorus (P) is a critical limiting nutrient in agroecosystems requiring careful management to reduce transport risk to aquatic environments. Routine laboratory measures of P bioavailability are based on chemical extractions performed on dried samples under oxidizing conditions. While useful, these tests are limited with respect to characterizing P release under prolonged water saturation. Labile orthophosphate bound to oxidized iron and other metals can rapidly desorb to solution in reducing environments, increasing P mobilization risk to surface runoff and groundwater. To better quantify P desorption potential and mobility during extended saturation, a laboratory microcosm method was developed based on repeated sampling of porewater and overlying floodwater over time. The method is useful for quantifying P release potential from soils and sediments varying in physicochemical properties and can improve site-specific P mitigation efforts by better characterizing P release risk in hydrologically active areas. Advantages of the method include its ability to simulate in situ dynamics, simplicity, low cost, and flexibility.

Accurate quantification of phosphorus (P) desorption potential in saturated soils and sediments is important for P modeling and transport mitigation efforts. To better account for in situ soil-water redox dynamics and P mobilization under prolonged saturation, a simple approach was developed based on repeated sampling of laboratory microcosms.

Phosphorus (P) is a critical limiting nutrient in agroecosystems requiring careful management to reduce transport risk to aquatic environments. Routine ...

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

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

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

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

In Situ Soil Moisture Sensors in Undisturbed Soils

Soil moisture directly affects operational hydrology, food security, ecosystem services, and the climate system. However, the adoption of soil moisture data has been slow due to inconsistent data collection, poor standardization, and typically short record duration. Soil moisture, or quantitatively volumetric soil water content (SWC), is measured using buried, in situ sensors that infer SWC from an electromagnetic response. This signal can vary considerably with local site conditions such as clay content and mineralogy, soil salinity or bulk electrical conductivity, and soil temperature; each of these can have varying impacts depending on the sensor technology.
Furthermore, poor soil contact and sensor degradation can affect the quality of these readings over time. Unlike more traditional environmental sensors, there are no accepted standards, maintenance practices, or quality controls for SWC data. As such, SWC is a challenging measurement for many environmental monitoring networks to implement. Here, we attempt to establish a community-based standard of practice for in situ SWC sensors so that future research and applications have consistent guidance on site selection, sensor installation, data interpretation, and long-term maintenance of monitoring stations.
The videography focuses on a multi-agency consensus of best-practices and recommendations for the installation of in situ SWC sensors. This paper presents an overview of this protocol along with the various steps essential for high-quality and long-term SWC data collection. This protocol will be of use to scientists and engineers hoping to deploy a single station or an entire network.

In Situ Soil Moisture Sensors in Undisturbed Soils

The determination of soil water content is a critical mission requirement for many state and federal agencies. This protocol synthesizes multi-agency efforts to measure soil water content using buried in situ sensors.

Soil moisture directly affects operational hydrology, food security, ecosystem services, and the climate system. However, the adoption of soil moisture ...

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface in response to disturbances. Deciphering the rapid biogeochemical changes requires in situ, minimally invasive tools with high spatial and temporal sampling resolution. However, the available passive sampling devices are not very useful in many cases either due to their disposable nature or the complexity and extensive workload for sample preparation.
To address this problem, a microdialysis profiler with 33 individual polyethersulfone nanomembrane tubes (semipermeable, &#60;20 nm pore size) integrated into the one-dimensional skeleton (60 mm) was established to iteratively sample the dissolved compounds in porewater across the soil-water interface at a high resolution of 1.8 mm (outer diameter plus one spacing, i.e., 0.1 mm between probes). The sampling mechanism is based on the principle of concentration gradient diffusion. The automatic loading of degassed water allows minimal disturbance to the chemical species across the oxic-anoxic interface.
This paper describes the procedures of device setup and continuous porewater sampling across the soil-water interface on a daily basis. Concentration-depth profiles were selectively measured before (on Day 6) and after (on Day 7) disturbances induced by irrigation. The results showed that concentration-depth profiles were undergoing rapid changes, especially for redox-sensitive elements (i.e., iron and arsenic). These protocols can help investigate the biogeochemical responses across the soil-water interface under various disturbances caused by physical, chemical, and biological factors. The paper thoroughly discusses the advantages and disadvantages of this method for potential use in the environmental sciences.

A microdialysis profiler is described to sample dissolved porewater solutes across an oxic-anoxic soil-water interface in situ with minimal disturbance. This device is designed to capture rapid changes in concentration-depth profiles induced by disturbances at the soil-water interface and beyond.

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface ...

Determination of Moisture Content in Soil

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba - The University of Arizona 
Demonstrating Author: Bradley Schmitz
Soils normally contain a finite amount of water, which can be expressed as the &#8220;soil moisture content.&#8221; This moisture exists within the pore spaces in between soil aggregates (inter-aggregate pore space) and within soil aggregates (intra-aggregate pore space) (Figure 1). Normally this pore space is occupied by air and/or water. If all the pores are occupied by air, the soil is completely dry. If all the pores are filled with water, the soil is said to be saturated.
<img alt="Figure 1" src="/files/ftp_upload/10011/10011fig1.jpg" /> 
Figure 1. Pore space in soil.

Source: Laboratories of Dr. Ian Pepper and Dr. Charles Gerba - The University of Arizona
Demonstrating Author: Bradley Schmitz
Soils normally contain a ...

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

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