The authors wish to thank Kalyani Muhunthan for assistance in laboratory procedures, Lee Opdahl for wheat genotype harvesting, the Washington State Concord Grape Research Council, and the Washington State University Center for Sustaining Agriculture and Natural Resources for a BIOAg grant to support this work. Additional funding was provided by the USDA/NIFA through Hatch project 1014527.

NOTE: Location of Field Site: Washington State University, Plant Pathology Farm (46&#176;46&#8217;38.0&#8221;N 117&#176;04&#8217;57.4&#8221;W). Seeds were sown using a mechanical planter on October 19, 2017. Each wheat genotype was planted in headrows, approximately 1 meter apart to avoid overlapping of root system. Plant and soil samples were collected on August 9, 2018, when plants were ready for harvest. Samples were gathered from three replicates of four wheat genotypes: PI561727, PI561725, Madsen, Lewjain.
1. Preparation of modified M9 medium
<ol>
	<li>Use Na2PO4&#8729;7H2O (12.8 g/200 mL), KH2PO4 (0.3 g/200 mL), NaCl (0.5 g/200 mL), and NH4Cl (1 g/20 mL) reagents to prepare the M9 salt solution.</li>
	<li>Use 18 g of MgSO4&#8729;7H2O in 100 mL of double deionized water (ddH2O) to prepare 0.75 M MgSO4&#8729;7H2O.</li>
	<li>Make 1 M of CaCl2&#8729;2H2O by adding 14.7 g of CaCl2&#8729;2H2O with 100 mL of ddH2O.</li>
	<li>Prepare the buffer solution by dissolving 6.048 g of PIPES into 156 mL of ddH2O with stirring. Adjust the pH to 6.8 with 5 M NaOH.</li>
	<li>Prepare 20% glucose (dextrose monohydrate) by dissolving 20 g of glucose into 100 mL of ddH2O.</li>
	<li>Prepare solutions and individually autoclave all but the glucose. Filter sterilize the glucose solution for addition to the M9 media after autoclaving (0.22 &#181;m).</li>
	<li>Prepare the modified M9 medium by mixing 156 mL of PIPES Buffer solution, 40 mL of M9 salts solution, 20 &#956;L of CaCl2&#183;2H2O solution, 266 &#956;L of MgSO4&#183;7H2O, and 4 mL of 20% glucose solutions together in the biosafety cabinet, using sterile technique.</li>
	<li>For preservation of the modified M9, seal the container, cover with aluminum foil to protect from UV, and place at 4 &#176;C.</li>
</ol>
2. Preparation of CAS-Fe-Agar medium
<ol>
	<li>Acid wash all glassware in 100 mM HCl/100 mM HNO3 for a minimum of 2 h prior to utilization in the CAS assay.</li>
	<li>Prepare an aluminum baking pan filled with laboratory grade sand and cover it with aluminum foil. Autoclave at 121 &#176;C for 30 min and set aside.</li>
	<li>Prepare HDTMA (hexadecyltrimethylammonium bromide) by adding 0.0365 g to 20 mL ddH2O and place the solution at 37 &#176;C to promote solubilization.</li>
	<li>Prepare 10 mM HCl and generate 1 mM FeCl3&#183;6H2O using 10 mM HCl as the solvent. Add 0.0302 g of CAS to 25 mL of ddH2O while gently stirring with a sterile magnetic stir bar. Then add 5 mL of 1 mM FeCl3&#183;6H2O (in 10 mM HCl) to the 25 mL of CAS solution while continuing to gently stir (the solution turns into dark reddish black color).</li>
	<li>Slowly add the 20 mL of HDTMA solution while gently stirring, into the Fe-CAS solution (this yields a dark blue solution).</li>
	<li>Prepare the buffer solution by dissolving 15.12 g of PIPES into 375 mL of ddH2O, with gentle stirring. Adjust the pH to 6.8 with 5 M NaOH. Add water to bring the volume to 450 mL. Add 5 g of agarose to the solution. Autoclave the PIPES buffer solution and the CAS-Fe solution at 121 &#176;C for 30 min.</li>
	<li>Carefully add the entirety of the CAS-Fe solution to the entirety of the PIPES Buffer in the biosafety cabinet after each of them is autoclaved.</li>
	<li>Place the mixed solution in a water bath at 50 &#176;C. 
	NOTE: Freshly prepare all the reagents in the CAS-Fe-Agar medium before each assay, as long-term storage at 50 &#176;C results in precipitation of the CAS-Fe complex, and cooling results in solidified medium.</li>
	<li>Place a sterile reagent boat in the sterile sand in the biosafety cabinet and heat to 50 &#176;C. Transfer the CAS-Fe-Agar to the boat, then quickly aliquot 100 &#181;L to each well in a clear, flat-bottom, sterile 96-well microplate.</li>
</ol>
3. Pyoverdine/EDTA standard preparation
<ol>
	<li>Pyoverdine preparation
	<ol>
		<li>Prepare 800 &#956;M pyoverdine standard (mixture of succinic acid, 2-hydroxy glutaramide, and succinaminde forms of pyoverdine &#8211; see Table of Materials), in previously prepared modified M9 medium.</li>
		<li>Dilute this solution into 400, 200, 100, 50, 25, 12.5, and 6.25 &#956;M solutions.</li>
	</ol>
	</li>
	<li>EDTA standard preparation
	<ol>
		<li>Add 0.594 g of disodium ethylenediaminetetraacetic acid (EDTA: C10H14N2Na2O8.2H2O), to 500 mL of previously prepared modified M9 medium to prepare 3200 &#956;M EDTA standard.</li>
		<li>Dilute this solution into 1600, 800, 400, 200, 100, 50, 25, 12.5, and 6.25 &#956;M solutions.</li>
	</ol>
	</li>
	<li>Standard curve generation
	<ol>
		<li>Add 100 &#956;L of each concentration of pyoverdine and EDTA to separate wells of a 96-well microplate containing 100 &#956;L of CAS-Fe Agar medium. Make duplicate technical replications of each concentration. Also, add blank wells with only M9 (no EDTA or pyoverdine).</li>
		<li>Using a microplate reader, measure absorbance (at 420 nm and 665 nm) after 1, 6, and 24 h incubation at 22 &#176;C, and use absorbance measurements to generate standard curves. 
		NOTE: For 420 nm measurements, subtract absorbance of blanks from the absorbance of pyoverdine or EDTA containing wells. For 665 nm, subtract absorbance of pyoverdine or EDTA containing wells from absorbance of blanks. Then, log10(&#181;M pyoverdine or EDTA) is regressed against the absorbance measurements. For ease of interpreting sample results, use absorbance as the x-axis and log10(&#181;M pyoverdine or EDTA) as the y-axis.</li>
	</ol>
	</li>
</ol>
4. Collection of environmental samples: soil and plant tissues
<ol>
	<li>Wash sampling equipment (shovels and scissors) with 0.22 &#181;m filtered ddH2O followed by 70% ethanol, and wipe with paper towels before sampling and in between samples to maintain sterile technique and reduce cross-contamination.</li>
	<li>Excise the plant tissues (grains and shoots) from plants in the field, and place them in a labeled plastic storage bag, leaving enough shoot stubble for ease in uprooting the desired soil and root samples.</li>
	<li>Excavate a small root ball approximately 15 cm deep and 23 cm wide and place it in a separate, labeled plastic bag for sample preparation in the laboratory environment. This step is similar to the methods of McPherson et al.27.</li>
	<li>Place all samples (grains, shoots, bulk soil, and root balls in separate bags) directly on ice and keep at 4 &#176;C until samples are processed for the siderophore production assay.</li>
	<li>Separate root associated soil samples into bulk, loosely-bound rhizosphere soil, and tightly-bound rhizosphere soil.
	<ol>
		<li>Take the root balls out of the bags. Gently shake off soil from the root ball. Shaken off soil, along with the soil left in the bag comprises the &#8220;bulk&#8221; soil.</li>
		<li>Use a rubber mallet to further remove soil from the root ball. This is the loosely-bound rhizosphere soil.</li>
		<li>Generate tightly-bound rhizosphere soil by taking roots with the tightly-bound sample and putting them in a centrifuge tube. Add 30 mL of ddH2O and vortex it for 2-3 minutes. Remove roots to get the tightly-bound rhizosphere soil slurry dilution.</li>
	</ol>
	</li>
</ol>
5. Preparation of siderophore enrichment cultures and CAS-Fe siderophore production assay
NOTE: All glassware should be acid washed prior to beginning the assays.
<ol>
	<li>Soil sample preparation (for each of the three soil sample types)
	<ol>
		<li>Homogenize each soil sample within the sample bag, by mixing and turning the soil as much as possible without opening the bag. 
		NOTE: This helps reduce natural soil spatial variability and aligns with normal soil sampling procedures. Other methods may be utilized to homogenize environmental samples, as appropriate, and depending on the experimental design.</li>
		<li>After each sample has been thoroughly mixed, aliquot and suspend 2.0 g of each soil in 20 mL of modified M9 medium, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
		<li>For tightly-bound rhizosphere samples, add 2 mL of the rhizosphere soil slurry to 20 mL of modified M9 medium, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
	</ol>
	</li>
	<li>Tissue sample (root, shoot and grain) preparation
	<ol>
		<li>Surface sterilize the sample with 70% ethanol. Macerate 2.0 g of fresh tissue in 20 mL of modified M9 medium using a blender on high for 30 seconds. Transfer the sample to a sterile 50 mL centrifuge tube, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
	</ol>
	</li>
	<li>Enrichment of Siderophore Production Through Fe Limitation
	<ol>
		<li>Incubate 50 mL centrifuge tubes at room temperature and shake at 160 rpm.</li>
	</ol>
	</li>
</ol>
6. CAS-Fe agar assays for detection of siderophore production in environmental samples
<ol>
	<li>At 24, 48, and 72 h after initiating the enrichment culture, remove 1 mL subsamples from the enrichment tubes using sterile technique and centrifuge at 10,000 x g for 1 min in 2 mL centrifuge tubes to pelletize the cells.</li>
	<li>Collect the separated supernatant. Using sterile technique, add 100 &#956;L of the supernatant to 100 &#956;L solution of CAS-Fe-Agar in duplicate or triplicate in the microplate. Also add 100 &#181;L of sterile M9 medium (as blanks). Then incubate the plate at 28 &#176;C.</li>
	<li>Suspend the remaining supernatant and pellet for each sample (not added to microtiter plate) into its own, sterile 2 mL centrifuge tube. Add 400 &#956;L of sterile glycerol into each sample-supernatant tube and resuspend the pellet to create glycerol stocks. Freeze the stock at -80 &#176;C for later analyses. 
	NOTE: This step can be modified to generate glycerol stocks according to any preferred in-house protocol.</li>
	<li>Measure absorbance at 6, 24, 48, and 72 h, at 420 nm wavelength.</li>
	<li>Use standard curves generated from pyoverdine or EDTA to interpret sample absorbance measurements in terms of pyoverdine equivalents. 
	NOTE: Pyoverdine was determined to be a superior standard compared with EDTA (vide infra), so EDTA was not used to interpret results in the current study.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The primary result of this work is the production of a new methodology that can be used to rapidly enrich for siderophore producing microbes while quantitatively measuring siderophore production/activity in the environmental sample. The methodology is quick, simple, and cost-effective, and the results show how it can be used to detect siderophore activity from complex and novel sample types (e.g., soil and plant tissue). The protocol also results in the production of glycerol stocks of the enrichment cultures, w...

Siderophores are important biomolecules involved primarily in iron-chelation for bioavailability, but with a wide array of additional purposes in terrestrial and aquatic ecosystems ranging from microbial quorum sensing, signaling to microbial plant-hosts, plant growth promotion, cooperation and competition within complex microbial communities1,2. Siderophores can be broadly classified according to their active sites and structural features, creating four basic types: carboxylate, hydroxamate, catecholate, and mixed types3,4. Many microorganisms are capable of excreting more than one type of siderophore5 and in complex communities, a vast majority of organisms biosynthesize the membrane receptors to allow the uptake of an even wider variety of siderophores1,6. Recent work indicates that siderophores are particularly important at the community level, and even in inter-kingdom communications and biogeochemical transfers7,8,9,10,11.
Chrome azurol sulphonate (CAS) has been used for over 30 years as a chelating agent to bind iron (Fe) in such a way that addition of ligands (i.e., siderophores) can result in dissociation of the CAS-Fe complex, creating an easily identifiable color change in the medium12. When the CAS is bound with Fe, the dye appears as a royal blue color, and as the CAS-Fe complex dissociates, the medium changes color according to the type of ligand used to scavenge the Fe13. The initial, liquid-based medium established by Schwyn and Neilands in 1987, has been modified in many ways to accommodate changing microbial targets14, growth habits and limitations15, as well as a variety of metals besides Fe, including aluminum, manganese, cobalt, cadmium nickel, lithium, zinc16, copper17, and even arsenic18.
Many human pathogens, as well as plant growth promoting microorganisms (PGPM) have been identified as siderophore-producing organisms3,19,20, and important rhizosphere and endophytic PGPM often test positive for siderophore-production4. The traditional Fe-based liquid method has been adapted to microtiter testing of isolates in cultivation for siderophore production21. However, these techniques fail to recognize the importance of the microbial community as a whole (the microbiome), in cooperation and potential regulation of siderophore production in soils and plant systems22. For that reason, we have developed a high-throughput community-level assessment of siderophore production from a given environment, based on the traditional CAS assay, but with replication, ease of measurement, reliability, and repeatability in a microplate assay.
In this study, a cost-effective, high-throughput CAS-Fe assay for detecting siderophore production is presented to assess the enrichment of siderophore production from complex samples (i.e., soil and plant tissue homogenates). Bulk, loosely-bound, and tightly-bound rhizosphere soil (in terms of how the soil was bound to the root) were obtained along with grain, shoot, and root tissues from four distinct wheat (Triticum aestivum L.) genotypes: Lewjain, Madsen, PI561725, and PI561727. It was hypothesized that fundamental differences in the wheat genotypes could result in differences in recruitment and selection of siderophore producing communities. Of particular interest is the difference between microbial communities associated with the PI561725 isogenic line, which is aluminum tolerant because it possesses ALMT1 (Aluminum-activated Malate Transporter 1), compared with the aluminum sensitive PI561727 isogenic line, which possesses a non-aluminum responsive form of the gene, almt123,24,25,26. The chief objective of the study was to develop a straightforward, rapid method of quantitatively assessing siderophore production in siderophore enrichment cultures of complex sample types while preserving the cultures for future work.

<table>
	<tbody>
		<tr>
			<td>Agarose</td>
			<td>Apex</td>
			<td>LF451320014</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum Baking Pan</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum Foil</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ammonium chloride, granular</td>
			<td>Fiesher Scientific</td>
			<td>152315A</td>
			<td></td>
		</tr>
		<tr>
			<td>Autoclave and Sterilizer</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Calcium chloride dihydrate</td>
			<td>Fiesher Scientific</td>
			<td>171428</td>
			<td></td>
		</tr>
		<tr>
			<td>CAS (Chrome Azurol S)</td>
			<td>Chem-Impex Int&#39;l Inc)</td>
			<td>000331-27168</td>
			<td></td>
		</tr>
		<tr>
			<td>Dextrose Monohydrate (glucose), crystalline powder</td>
			<td>Fiesher Scientific</td>
			<td>1521754</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA, disodium salt, dihydrate, Crystal</td>
			<td>J.T.Baker</td>
			<td>JI2476</td>
			<td></td>
		</tr>
		<tr>
			<td>Glycerol, Anhydrous</td>
			<td>Baker Analyzed</td>
			<td>C22634</td>
			<td></td>
		</tr>
		<tr>
			<td>HDTMA (Cetyltrimethylammomonium Bromide</td>
			<td>Reagent World</td>
			<td>FZ0941</td>
			<td></td>
		</tr>
		<tr>
			<td>Hydrochloride acid</td>
			<td>ACROS Organic</td>
			<td>B0756767</td>
			<td></td>
		</tr>
		<tr>
			<td>Infinite M200 PRO plate reader</td>
			<td>TECAN</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Iron (III) chloride hexahydrate, 99%</td>
			<td>ACROS Organic</td>
			<td>A0342179</td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory Fume Hood</td>
			<td>Thermo Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Laboratory Incubator</td>
			<td>VWR Scientific</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Magnesium Sulfate</td>
			<td>Fiesher Scientific</td>
			<td>27855</td>
			<td></td>
		</tr>
		<tr>
			<td>Niric Acid, (69-70)%</td>
			<td>J.T.Baker</td>
			<td>72287</td>
			<td></td>
		</tr>
		<tr>
			<td>PIPES buffer, 98.5%</td>
			<td>ACROS Organic</td>
			<td>A0338723</td>
			<td></td>
		</tr>
		<tr>
			<td>Potassium phosphate, dibaisc,powder</td>
			<td>J.T.Baker</td>
			<td>J48594</td>
			<td></td>
		</tr>
		<tr>
			<td>Pyoverdine</td>
			<td>SIGMA-ALDRICH</td>
			<td>078M4094V</td>
			<td></td>
		</tr>
		<tr>
			<td>Sand</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>SI-600R Shaker</td>
			<td>Lab Companion</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride, granular</td>
			<td>Fiesher Scientific</td>
			<td>136539</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide, pellets</td>
			<td>J.T.Baker</td>
			<td>G48K53</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium phosphate, dibasic heptahydrate, 99%</td>
			<td>ACROS Organic</td>
			<td>A0371705</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

A pyoverdine mixture biosynthesized by Pseudomonas fluorescens was used as a standard to interpret and quantify absorbance (at 420 nm) of samples in terms of pyoverdine equivalents in &#181;M. Figure 1 shows the relationship between absorbance (420 nm) and starting concentration of pyoverdine (Log10 molarity in &#181;M). EDTA did not provide an adequate standard because samples exhibited greater absorbance measurements than were attainable...

Watch this Scientific Journal Video about High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils at JoVE.com

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

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 ...

high-throughput-siderophore-screening-from-environmental-samples

Research

JoVE Journal

Environment

12.5K Views.  Washington State University. We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

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

Determination of Microbial Extracellular Enzyme Activity in Waters, Soils, and Sediments using High Throughput Microplate Assays

Much of the nutrient cycling and carbon processing in natural environments occurs through the activity of extracellular enzymes released by microorganisms. Thus, measurement of the activity of these extracellular enzymes can give insights into the rates of ecosystem level processes, such as organic matter decomposition or nitrogen and phosphorus mineralization. Assays of extracellular enzyme activity in environmental samples typically involve exposing the samples to artificial colorimetric or fluorometric substrates and tracking the rate of substrate hydrolysis. Here we describe microplate based methods for these procedures that allow the analysis of large numbers of samples within a short time frame. Samples are allowed to react with artificial substrates within 96-well microplates or deep well microplate blocks, and enzyme activity is subsequently determined by absorption or fluorescence of the resulting end product using a typical microplate reader or fluorometer. Such high throughput procedures not only facilitate comparisons between spatially separate sites or ecosystems, but also substantially reduce the cost of such assays by reducing overall reagent volumes needed per sample.

Microplate based procedures are described for the colorimetric or fluorometric analysis of extracellular enzyme activity. These procedures allow for the rapid assay of such activity in large numbers of environmental samples within a manageable time frame.

Much of the nutrient cycling and carbon  processing in natural environments occurs through the activity of extracellular  enzymes released by ...

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.

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 ...

Measurement of Greenhouse Gas Flux from Agricultural Soils Using Static Chambers

Measurement of greenhouse gas (GHG) fluxes between the soil and the atmosphere, in both managed and unmanaged ecosystems, is critical to understanding the biogeochemical drivers of climate change and to the development and evaluation of GHG mitigation strategies based on modulation of landscape management practices. The static chamber-based method described here is based on trapping gases emitted from the soil surface within a chamber and collecting samples from the chamber headspace at regular intervals for analysis by gas chromatography. Change in gas concentration over time is used to calculate flux. This method can be utilized to measure landscape-based flux of carbon dioxide, nitrous oxide, and methane, and to estimate differences between treatments or explore system dynamics over seasons or years. Infrastructure requirements are modest, but a comprehensive experimental design is essential. This method is easily deployed in the field, conforms to established guidelines, and produces data suitable to large-scale GHG emissions studies.

This article showcases the static chamber-based method for measurement of greenhouse gas flux from soil systems. With relatively modest infrastructure investments, measurements may be obtained from multiple treatments/locations and over timeframes ranging from hours to years.

Measurement of greenhouse gas (GHG) fluxes between the soil and the atmosphere, in both managed and unmanaged ecosystems, is critical to understanding the ...

Automated Gel Size Selection to Improve the Quality of Next-generation Sequencing Libraries Prepared from Environmental Water Samples

Next-generation sequencing of environmental samples can be challenging because of the variable DNA quantity and quality in these samples. High quality DNA libraries are needed for optimal results from next-generation sequencing. Environmental samples such as water may have low quality and quantities of DNA as well as contaminants that co-precipitate with DNA. The mechanical and enzymatic processes involved in extraction and library preparation may further damage the DNA. Gel size selection enables purification and recovery of DNA fragments of a defined size for sequencing applications. Nevertheless, this task is one of the most time-consuming steps in the DNA library preparation workflow. The protocol described here enables complete automation of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments. 
In this study, we describe a high-throughput approach to prepare high quality DNA libraries from freshwater samples that can be applied also to other environmental samples. We used an indirect approach to concentrate bacterial cells from environmental freshwater samples; DNA was extracted using a commercially available DNA extraction kit, and DNA libraries were prepared using a commercial transposon-based protocol. DNA fragments of 500 to 800 bp were gel size selected using Ranger Technology, an automated electrophoresis workstation. Sequencing of the size-selected DNA libraries demonstrated significant improvements to read length and quality of the sequencing reads.

This manuscript describes an automated gel size selection approach for purifying DNA fragments for next-generation sequencing. The Ranger Technology provides complete automation of the entire process of agarose gel loading, electrophoretic analysis, and recovery of targeted DNA fragments allowing for high-throughput and high quality next-generation sequencing libraries.

Next-generation sequencing of environmental samples can be challenging because of the variable DNA quantity and quality in these samples. High quality DNA ...

Physical Isolation of Endospores from Environmental Samples by Targeted Lysis of Vegetative Cells

Endospore formation is a survival strategy found among some bacteria from the phylum Firmicutes. During endospore formation, these bacteria enter a morpho-physiological resting state that enhances survival under adverse environmental conditions. Even though endospore-forming Firmicutes are one of the most frequently enriched and isolated bacterial groups in culturing studies, they are often absent from diversity studies based on molecular methods. The resistance of the spore core is considered one of the factors limiting the recovery of DNA from endospores. We developed a method that takes advantage of the higher resistance of endospores to separate them from other cells in a complex microbial community using physical, enzymatic and chemical lysis methods. The endospore-only preparation thus obtained can be used for re-culturing or to perform downstream analysis such as tailored DNA extraction optimized for endospores and subsequent DNA sequencing. This method, applied to sediment samples, has allowed the enrichment of endospores and after sequencing, has revealed a large diversity of endospore-formers in freshwater lake sediments. We expect that the application of this method to other samples will yield a similar outcome.

A method to single out bacterial endospores from complex microbial communities was developed to perform tailored culture or molecular studies of this group of bacteria.

Endospore formation is a survival strategy found among some bacteria from the phylum Firmicutes. During endospore formation, these bacteria enter a ...

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 ...

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 ...

A 3-dimensional (3D)-printed Template for High Throughput Zebrafish Embryo Arraying

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related research fields. Thanks to its unique features, including large fecundity, embryo translucency, rapid and simultaneous development, etc., zebrafish embryos are often used for large scale toxicity assessment of chemicals and drug/compound screening. A typical screening procedure involves adult zebrafish spawning, embryos selection, and arraying the embryos into multi-well plates. From there, embryos are subjected to exposure and the toxicity of chemical, or the effectiveness of the drugs/compounds can be evaluated relatively quickly based on phenotypic observations. Among these processes, embryos arraying is one of the most time-consuming and labor-intensive steps that limits the throughput level. In this protocol, we present an innovative approach that makes use of a 3D-printed arraying template coupled with vacuum manipulation to speed up this laborious step. The protocol herein describes the overall design of the arraying template, a detailed experimental setup and step-by-step procedure, followed by representative results. When implemented, this approach should prove beneficial in a variety of research applications using zebrafish embryos as testing subjects.

A 3-dimensional 3D-printed Template for High Throughput Zebrafish Embryo Arraying

Here, we present a protocol to design and fabricate a zebrafish embryo arraying template, followed by a detailed procedure on the use of such template for high throughput zebrafish embryo arraying into a 96-well plate.

The zebrafish is a globally recognized fresh water organism frequently used in developmental biology, environmental toxicology, and human disease related ...

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.