This work was supported by startup funding from the University of Utah Department of Chemistry and NSF CAREER Award #2339190. We thank members of the Puri Lab for helpful discussions. We thank Rachel Hurrell (University of Utah) for initial guidance with the flow cytometry experiment.

The details of the reagents and the equipment used in the study are listed in the Table of Materials.
1. Preparation and extrusion of gradient syringes
NOTE: Gradient syringe preparation should be performed using&#160;sterile technique.
<ol>
	<li>Use several colonies of Methylomonas sp. LW13 freshly grown on a plate to inoculate 6 mL of nitrate mineral salts (NMS) medium in an 18 mm x 150 mm glass tube. Seal the tube with a serum stopper and aluminum crimp seal, and add methane using a syringe to a final atmosphere of 50% (v/v) methane in&#160;air. Shake this planktonic liquid culture at 200 rpm at room temperature until turbid (about one day).</li>
	<li>Passage liquid cultures 1:10 into fresh media. Continue growing liquid cultures of methanotrophs to log-phase growth (OD600 of ~0.5) and adjust to an OD600 =1.0.</li>
	<li>Prepare syringes by removing the accompanying plunger and keeping them in a sterile container. Attach a sterile PTFE filter tip to the syringe and place it in a standard test tube rack with the tip facing down.</li>
	<li>For each 10 mL syringe, thoroughly mix 1 mL of cells from step 1.2 with 5 mL of NMS and 4 mL of molten agarose (0.5% m/v, cooled to 55 &#176;C) in a sterile conical tube. These volumes can be scaled up to fill multiple syringes in parallel.</li>
	<li>Slowly pour or use a serological pipet to add the mixture to each syringe, up to the 8 mL marking. Allow agarose within syringes to solidify (~15 min), then cap with a sterile 20 mm rubber butyl stopper. Secure the stopper to the syringe using lab tape and label it with syringe contents.</li>
	<li>To add methane to the syringe headspace, fill a large (60 mL) syringe with 100% CH4 and attach a PTFE filter tip (0.2 &#181;m, 25 mm) connected to a sterile needle (23 G). Pierce the rubber stopper with the large syringe and pierce a second sterile needle through the stopper to create a gas outlet.</li>
	<li>Depress the plunger on the large syringe to allow 20 mL of 100% CH4 to flush through the headspace, taking care to remove the outlet needle when there is 1-2 mL of CH4 left in the large syringe to prevent oxygen backflow through the outlet needle.</li>
	<li>Incubate the syringes at 18 &#176;C, repeating steps 1.6 and 1.7 daily to replenish the methane.</li>
	<li>To extrude agarose, replace the PTFE filter tip with a sterile 23 G needle and replace the rubber stopper with the supplied syringe plunger. Slowly depress the plunger to dispense 1 mL increments into separate sterile 1.5 mL microcentrifuge tubes.</li>
</ol>
2. Determining counter gradient gas concentrations
<ol>
	<li>Measuring the dissolved oxygen gradient
	<ol>
		<li>Use a razor blade to slice through the width of an agarose-filled syringe (prepared following steps 1.1-1.8) close to the PTFE filter. Fasten the opened syringe to a syringe pump oriented towards a Clark-type microelectrode, with the open end facing the tip of the electrode.</li>
		<li>Adjust the settings of the syringe pump to move the syringe towards the microelectrode at a rate of 1 mL/min (0.6 cm/min); begin recording the dissolved oxygen measurements on the Unisense Logger software as soon as the syringe pump begins moving.</li>
	</ol>
	</li>
	<li>Measuring the methane gradient
	<ol>
		<li>Immediately before extrusion, replace the syringe filter tip with a one-way stopcock connected to a 23 G needle, and quickly swap the rubber stopper with a syringe plunger. Add eight 1 mL agarose aliquots to separate evacuated 12 mL gas-tight vials and allow the samples to equilibrate at room temperature for 1 h.</li>
		<li>Equilibrate sample vials to atmospheric pressure by cracking open and immediately resealing vials or piercing and quickly removing a needle. Inject 500 &#181;L of vial headspace into a gas chromatograph with flame ionization detection (GC-FID) using a gas-tight syringe. Create a calibration curve derived from CH4 standards to convert peak area (pA*min) to &#181;mol/L.</li>
	</ol>
	</li>
</ol>
3. Counting cells in the gradient syringe
<ol>
	<li>Flow cytometry
	<ol>
		<li>Extrude 1 mL of agarose segments from gradient syringes inoculated with either wild-type or mutant LW13 as outlined in step 1.9. Also, prepare and extrude agarose from a cell-free, sterile syringe as a negative control.</li>
		<li>Add 0.75 mL of 0.85% (m/v) NaCl in water to all extruded agarose samples and homogenize by vortexing. Further, dilute samples 1:10 by transferring 100 &#181;L into a new microcentrifuge tube and adding 900 &#181;L of the salt solution.</li>
		<li>Add 3 &#181;L of a 1:1 mixture of SYTO9 and propidium iodide stains, then incubate in the dark at room temperature for 15 min. To determine the cells per mL of agarose, sonicate the microsphere counting bead suspension in a water bath for 5 min. Then, add 10 &#181;L of the suspension to each sample before flow cytometry analysis.</li>
		<li>Analyze samples with a flow cytometer10,11 with the following parameters: triggering on green fluorescence, flow rate of 10 &#181;L/s, and particle analysis rate below 1,000 particles/s.
		<ol>
			<li>Compare SSC vs. FITC dot plots between cell-free control samples and inoculated agarose samples to draw &#34;bacterial event&#34; voltage gates that exclude background agarose particles. Additionally, draw voltage gates for the microsphere counting beads, which should be consistent between samples.</li>
		</ol>
		</li>
		<li>To determine the concentration of cells in each agarose segment within the gradient syringe, use the following equation, noting that the dilution factor for the above protocol is 17.7275 and that 10-6 mL is the volume of one microsphere bead. 
		<img alt="Equation 1" src="/files/ftp_upload/67191/67191eq01.jpg" style="margin: auto;" /></li>
	</ol>
	</li>
	<li>Counting colony-forming units within the gradient syringe
	<ol>
		<li>Extrude 1 mL of agarose segments into separate, sterile 2 mL microcentrifuge tubes, add 800 &#181;L of NMS, and vortex for 10 s to aid in pipetting.</li>
		<li>Prepare a sterile 96-well plate by adding 180 &#181;L of NMS to each well. Add 20 &#181;L of diluted agarose samples to each well in the first column and pipet to mix.</li>
		<li>Using a multi-channel pipette, serially dilute samples tenfold by transferring 20 &#181;L from the first row of wells into the second row of wells and pipetting 10 times to mix. Continue this process until the last row of the plate.</li>
		<li>Label square grid plates containing NMS agar or media of choice. Using a multi-channel pipette, spot 5 &#181;L from a column of the 96-well plate onto the agar plate. Depending on the size of the agar plate, multiple columns can be spotted on the same plate.</li>
		<li>Incubate plates under 40% methane in&#160;air and grow at 18 &#176;C. Count bacterial colonies after 2-3 days and determine the colony-forming units per milliliter (CFU/mL).</li>
	</ol>
	</li>
</ol>
4. Biomolecule detection assays
<ol>
	<li>Polysaccharide assay
	<ol>
		<li>Extrude 1 mL of agarose segments into separate 2 mL microcentrifuge tubes and mix with 1 mL of a 1% (m/v) Na2CO3 solution in water. Heat samples to 80 &#176;C for 30 min with vortexing every 5-10 min, followed by centrifugation at 4,000 x g at 4 &#176;C for 20 min.</li>
		<li>Collect the supernatant combined with three volumes of 100% ethanol and incubate at 20 &#176;C for at least 2 h (or overnight).</li>
		<li>Collect the ethanol-precipitated polysaccharides by centrifugation at 16,100 x g at 4 &#176;C for 30 min. Remove the supernatant and air-dry the pellet. Resuspend the pellet in 100 &#181;L of deionized water.</li>
		<li>Measure the relative polysaccharide content of each agarose segment using a phenol-sulfuric acid colorimetric assay12,13. Combine 50 &#181;L of the resuspended extract with 150 &#181;L of concentrated sulfuric acid and 30 &#181;L of 5% (v/v) phenol in water in a clear 96-well plate.</li>
		<li>Measure the absorbance at 490 nm using a microplate reader and calculate the relative polysaccharide content of each agarose segment as a percentage of the absorbance of the agarose segment closest to the PTFE filter.</li>
	</ol>
	</li>
	<li>Protein assay
	<ol>
		<li>Extrude agarose into separate 1.5 mL microcentrifuge tubes and transfer 100 &#181;L of each aliquot to glass test tubes.</li>
		<li>Determine the total protein concentration using the test tube protocol of a BCA protein assay kit. Prepare albumin BSA standards with agarose extruded from a sterile syringe as the diluent.</li>
	</ol>
	</li>
	<li>Extracellular DNA assay
	<ol>
		<li>Extrude agarose into separate 1.5 mL microcentrifuge tubes and transfer 20 &#181;L of each to 0.2 mL microcentrifuge tubes.</li>
		<li>Measure DNA concentrations using the commercially available 1x dsDNA high-sensitivity assay kit following the manufacturer&#39;s protocol.</li>
	</ol>
	</li>
</ol>
5. RNA extraction
<ol>
	<li>Prepare the extraction buffer by combining the following in 800 mL of RNase-free water: 2.0 g of CTAB, 2.0 g of polyvinylpyrrolidone (PVP 40), 81.8 g of NaCl, 100 mM of Tris-HCl (pH 8.0), and 20 mM of EDTA. Bring up the volume to 1 L and autoclave; store at 4 &#176;C.</li>
	<li>Aliquot the prepared extraction buffer and add 1% (v/v) final concentration of beta-mercaptoethanol just before use. Warm the buffer to 65 &#176;C using a water bath or heat block.</li>
	<li>Divide each gradient syringe into 1 mL sections by extruding agarose into separate RNase-free 2 mL microcentrifuge tubes following the procedure in step 1.9.</li>
	<li>Centrifuge samples at 21,000 x g, 4 &#176;C, for 15 min and discard the supernatant, keeping samples on ice.</li>
	<li>Add 600 &#181;L of pre-warmed extraction buffer to each pelleted 1 mL of extruded agarose. Add approximately 200 &#181;L of zirconia/silica beads and homogenize samples for 3 min at 30 Hz/s using a bead beater, pausing halfway to place samples on ice for 2 min.</li>
	<li>Centrifuge the samples at 15,000 x g, 4 &#176;C, for 2 min to reduce foaming. Extract samples by adding 600 &#181;L of chloroform: isoamyl alcohol (24:1) to tubes and vortexing for 10 s.</li>
	<li>Centrifuge the samples at 15,000 x g, 4 &#176;C, for 8 min. Carefully transfer the upper aqueous phase to a new RNase-free microcentrifuge tube and add 600 &#181;L of chloroform: isoamyl alcohol (24:1) to the transferred upper phase and vortex for 10 s.</li>
	<li>Centrifuge the samples at 15,000 x g, 4 &#176;C, for 8 min and transfer the new upper aqueous phase to a new RNase-free microcentrifuge tube. Add an equal volume of isopropanol to the transferred upper phase and incubate samples for several hours at -20 &#176;C. Optional: samples can be left at -20 &#176;C overnight.</li>
	<li>Collect RNA-containing precipitates by centrifuging samples at 16,100 x g, 4 &#176;C for 30 min.</li>
	<li>Discard the supernatant and wash the pellet with 300 &#181;L of cold 75% (v/v) ethanol made with RNase-free water and centrifuge at 16,100 x g, 4 &#176;C, for 5 min.</li>
	<li>Wash the pellets again following step 5.10.</li>
	<li>After removing the ethanol supernatant, let the pellets air dry for 15 min. Dissolve pellets in 100 &#181;L of RNase-free water.</li>
	<li>Treat samples with DNase I at 37 &#176;C for 30 min following the manufacturer&#39;s protocol.</li>
	<li>Inactivate DNase I by adding 300 &#181;L of acid phenol: chloroform: IAA (125:24:1, pH 4.5). Vortex for 10 s and incubate at room temperature for 5 min.</li>
	<li>Centrifuge at 16,100 x g, 4 &#176;C, for 5 min and keep the upper aqueous phase by transferring it to a new RNase-free microcentrifuge tube.</li>
	<li>Optional: Pool RNA from the same segment across all replicate syringes by combining the upper phases into a conical tube.</li>
	<li>Add 1 volume of isopropanol equal to the volume of the RNA-containing upper phase and add 7.5 M of LiCl to a final concentration of 0.8 M, inverting several times to mix.</li>
	<li>Incubate the samples for several hours at -20 &#176;C. Optional: samples can be left at -20 &#176;C overnight.</li>
	<li>Centrifuge the samples at 16,100 x g, 4 &#176;C, for 15 min and carefully discard the supernatant. Wash the RNA-containing pellet twice by adding cold 70% (v/v) ethanol in RNase-free water, centrifuging at 16,100 x g, 4 &#176;C, for 5 min, and removing the supernatant.</li>
	<li>Allow the pellets to air dry at room temperature for 10 min and resuspend in 50 &#181;L of RNase-free water. Store the RNA samples at -80 &#176;C.</li>
	<li>Optional, depending on the RNA quality: Samples can be re-purified using an RNA purification kit to remove small (&#60;200 nt) RNAs.</li>
	<li>To confirm RNA samples do not contain residual DNA, use 1 &#181;L of purified RNA as the template for PCR amplification using universal bacterial 16S rRNA gene primers 27F/1492R. Include two additional PCR reactions containing 1 &#181;L of either genomic DNA (diluted 1:100) or nuclease-free water to serve as the positive and negative controls, respectively.</li>
	<li>Run the products on a 1% agarose TBE gel using gel electrophoresis14. 
	NOTE: RNA samples with no DNA contamination should result in the absence of a band in the sample lane. If RNA samples show DNA contamination, reprocess the samples starting from step 5.13. RNA is ready for downstream analysis and can optionally be analyzed to quantify its integrity by measuring the RNA Integrity Number (RIN).</li>
</ol>

Modeling Methanotroph Niches – The Gradient Syringe Model Ecosystem

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The authors declare no conflicts of interest.

Methods for methanotroph cultivation 
Methanotrophs have been studied for decades to understand their physiology, their individual and community behavior in the natural environment, and their potential for methane mitigation in industrial applications. Throughout these studies, much of the research conducted has been performed using homogenous planktonic cultures where spatial context is lost. The gradient syringe model ecosystem was developed to replicate the methane-oxygen counter gradient characteristic of natural methanotroph habitats in the lab, allowing researchers to study methanotrophs growing in an environment that more closely resembles where these organisms evolved.
Over the past 30 years, researchers have recreated the methane-oxygen counter gradient in the lab using a variety of methods, often with the primary goal of isolating and classifying methanotrophs from mixed methane-oxidizing consortia. These methods can be divided into two approaches, both involving the use of opposing chambers of methane and oxygen: suspending relatively undisturbed soil on a membrane16,17,18, or inoculating small amounts of soil or pure bacterial culture into a minimal medium in agarose7,8,19. The gradient syringe method described here combines the syringe-based approach of Dedysh and coworkers9 with the cultivation of methanotrophs from previous work by Amaral and Knowles8, and Schink and coworkers7. The latter of these methods laid the foundation for cultivating methanotrophs in a methane-oxygen counter gradient and used a continuous flow of methane and oxygen on either side of the agarose plug. While this provides a more constant environment, this approach adds complexity to the experimental setup and necessitates dedicated gas sources.
In contrast, the gradient syringe described here relies on daily flushing of the syringe to provide fresh methane, a process that takes less than a minute per syringe, while providing continuous access to atmospheric oxygen through a sterile PTFE filter tip. This simpler method may enable wider adoption of this model ecosystem for studying methanotrophs in a spatially resolved context. The described protocol also details chemical and molecular-level analyses that can be performed directly on bacteria incubated in the semi-solid agarose. As a result, bacteria do not need to be excised and cultured outside the agarose matrix before analysis, preserving the gas gradient conditions at the time of sampling.
Remarks on the protocol 
Because the bacteria are cultured within a polypropylene volumetric syringe, researchers can use the accompanying syringe plunger to accurately and reproducibly segment the agarose plug while maintaining the spatial integrity of the agarose matrix that still remains in the syringe barrel. Without the air-tight design inherent to the syringe, agarose plugs would need to be removed from the syringe barrel and sliced, introducing uncertainty in the volume of agarose segments, and releasing unquantifiable amounts of methane and dissolved oxygen into the atmosphere. Agarose extrusion through a sterile needle simplifies sample preparation and helps homogenize extruded segments without shearing bacterial cells. This method allows researchers to divide each inoculated gradient syringe into at least eight agarose segments and perform parallel experiments on methanotrophs growing in a range of oxygen and methane concentrations.
In optimizing RNA extraction from high-polysaccharide content agarose, it was found that common reagents like guanidium thiocyanate and TRIzol led to agarose gelation, which obstructed purification columns and resisted pelleting by centrifugation. Low RNA yields and quality were also a concern as large polysaccharide molecules can trap nucleic acids while small polysaccharides can co-precipitate with RNA20. Instead, an extraction buffer containing the cationic surfactant CTAB was used, which solubilizes lipid membranes20; and NaCl, which prevents CTAB-nucleic acid complexes from forming and allows nucleic acids to precipitate but keeps polysaccharides in solution21. RNases were denatured by the inclusion of &#946;-mercaptoethanol in the CTAB buffer. For the RNA-seq experiment, an optional column-based purification step was included to exclude small (&#60;200 nucleotides) RNAs before library preparation.
Limitations and considerations 
While the NMS and agarose provide a minimal medium matrix for cultivating methanotrophic bacteria, the gradient syringe as described here only recreates the gas gradients of methanotrophic habitats, but not other gradients present in those environments such as trace metals22, salinity23, or other nutrients24. It is possible these gradients can be added to a similar system in the future. Additionally, the volume of the syringe (8 mL agarose) limits the total biomass per syringe, necessitating pooling multiple syringes for some analyses (as described in step 5.16). Although the handheld syringe conveniently segments the agarose into 1 mL aliquots, its size also limits the headspace to approximately 4 mL, limiting the amount of bulk methane that can be stored for the cultivated microbes. Since methane oxidation rates are proportional to the growth rate of aerobic methanotrophs25, daily replenishment of headspace methane is recommended. While this may still result in periods of methane limitation, these periods are reproducible in the laboratory and likely mimic situations found in natural environments.
While using the gradient syringe, the presence of the agarose polysaccharides necessitates some adjustments to the assays used to analyze methanotrophs grown in this system. For example, protocols requiring the transfer of small volumes of extruded agarose need multiple dilution steps with thorough homogenization between each dilution for accurate pipetting. Additionally, in cases such as the polysaccharide assay where the polysaccharides inherent to the agarose matrix will react with the sulfuric acid-phenol reagent, the inclusion of a sterile, cell-free agarose negative control is essential. Early attempts to mitigate these issues by including the agarose-hydrolyzing enzyme &#946;-agarase were unsuccessful and introduced an unknown variable to the biological experiments. The use of multiple technical replicates, thorough dilution, homogenization, and the inclusion of controls can be used to mitigate most of the challenges inherent to the agarose matrix.
Applications 
In addition to single-strain studies, the gradient syringe can support the co-culture of multiple strains, and soil can be used as the inoculum in place of pure bacterial culture. The simple design of the gradient syringe model ecosystem is amenable to the culture of other types of microorganisms that exist at the interface between anoxic and oxic environments by using a different gas substrate, such as H2 or CO, in place of methane. In summary, the use of a simple, spatially resolved model ecosystem allows researchers to study the unique physiology and metabolic adaptations of anoxic-oxic microorganisms and can be used to link genes with organismal phenotypes.

Microorganisms living at an anoxic-oxic interface often serve important ecological roles1. One example is aerobic methane-oxidizing bacteria (methanotrophs), which exist in counter gradients of methane and oxygen in soils and sediments2. These microorganisms possess unique metabolic and physiological characteristics that enable them to exploit the gas gradients present in their environments and have been the subject of ongoing research for decades3,4,5. Currently, most published research about methanotrophs and methane-oxidizing communities is based on work with homogenous planktonic cultures that often fail to capture the spatial and chemical gradients that are inherent to their natural microbial habitats. This limitation hinders our understanding of microbial physiology and our ability to link genomic information to phenotypic traits.
This protocol reports a simple, laboratory-based model ecosystem that creates reproducible conditions for studying both specific methanotrophs, such as Methylomonas sp. strain LW13, and methane-oxidizing communities directly from environmental soil samples. Importantly, cultivation in the gradient syringe results in counter gradient-specific phenotypes that are not present in homogenous planktonic cultures6, highlighting the system&#39;s ability to unveil new aspects of methanotroph physiology. Inspired by previously published model ecosystems7,8,9, the gradient syringe is a simplified method that can be used to collect chemical and molecular information from microorganisms cultured using this approach.
The reported procedures for genetic, chemical, and molecular analyses have been modified to work reliably on microbial cultures grown within a semi-solid agarose matrix. These procedures may also be useful for analyzing bacteria grown in other semi-solid agarose-based systems, such as those used for bacterial soft agar swimming assays. Adapting these analyses to spatially resolved contexts may open new avenues for studying microbial life in more ecologically relevant environments.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1% Gas mix analytical standard</td>
			<td>Supelco</td>
			<td>22561</td>
			<td>1% each component in nitrogen: carbon monoxide, carbon dioxide, hydrogen, methane and oxygen</td>
		</tr>
		<tr>
			<td>100% Methane</td>
			<td>Airgas</td>
			<td>ME CP300</td>
			<td>chemically pure grade</td>
		</tr>
		<tr>
			<td>15 ppm Gas mix analytical standard</td>
			<td>Supelco</td>
			<td>23470-U</td>
			<td>15 ppm each component in nitrogen: methane, ethane, ethylene, acetylene, propane, propylene, propyne, and n-butane</td>
		</tr>
		<tr>
			<td>1x Nitrate mineral salts&#160;</td>
			<td></td>
			<td>see CAS numbers below</td>
			<td>Dissolve the following in Mili-Q water and autoclave:&#160; 0.2 g/L MgSO4&#183;7H2O, 0.2 g/L CaCl2&#183;6H2O, 1 g/L KNO3, and 30 &#956;M LaCl3. Before use, add trace elements to a 1X final concentration and phosphate buffer (pH 6.8) to a final concentration of 5.8 mM.</td>
		</tr>
		<tr>
			<td>23 G needle</td>
			<td>BD Biosciences</td>
			<td>305194</td>
			<td>sterile, Luer-Lok</td>
		</tr>
		<tr>
			<td>500x Trace elements</td>
			<td></td>
			<td>see CAS numbers below</td>
			<td>Dissolve the following in Milli-Q water: 1.0 g/L Na2-EDTA, 2.0 g/L FeSO4&#183;7H2O, 0.8 g/L ZnSO4&#183;7H2O, 0.03 g/L MnCl2&#183;4H2O, 0.03 g/L H3BO3, 0.2 g/L CoCl2&#183;6H2O, 0.6 g/L CuCl2&#183;2H2O, 0.02 g/L NiCl2&#183;6H2O, and 0.05 g/L Na2MoO&#183;2H2O.</td>
		</tr>
		<tr>
			<td>96 Well plate</td>
			<td>CELLTREAT</td>
			<td>229596</td>
			<td>sterile</td>
		</tr>
		<tr>
			<td>Acid phenol:chloroform:IAA (125:24:1)</td>
			<td>Invitrogen</td>
			<td>AM9720</td>
			<td>&#160;pH 4.5</td>
		</tr>
		<tr>
			<td>Agarose</td>
			<td>Fisher Scientific</td>
			<td>BP160</td>
			<td>molecular biology grade, CAS 9012-36-6</td>
		</tr>
		<tr>
			<td>Aluminum crimp seals</td>
			<td>VWR</td>
			<td>30618-460</td>
			<td>20 mm</td>
		</tr>
		<tr>
			<td>Bead beater</td>
			<td>Qiagen</td>
			<td>9003240</td>
			<td>TissueLyser III</td>
		</tr>
		<tr>
			<td>Butyl rubber stopper</td>
			<td>Chemglass Life Science</td>
			<td>50-143-854</td>
			<td>20 mm, blue</td>
		</tr>
		<tr>
			<td>Chloroform:isoamyl alcohol (24:1)</td>
			<td>Millipore Sigma</td>
			<td>25666</td>
			<td>BioUltra, for molecular biology</td>
		</tr>
		<tr>
			<td>Clark-type O2 microelectrode</td>
			<td>Unisense</td>
			<td>OX-500</td>
			<td></td>
		</tr>
		<tr>
			<td>DEPC-treated water</td>
			<td>Thermo Scientific</td>
			<td>R0601</td>
			<td></td>
		</tr>
		<tr>
			<td>DNase I (Ambion)</td>
			<td>Invitrogen</td>
			<td>AM2222</td>
			<td></td>
		</tr>
		<tr>
			<td>Flow cytometer</td>
			<td>Beckman Coulter</td>
			<td></td>
			<td>CytoFLEX</td>
		</tr>
		<tr>
			<td>Gas chromatograph (flame ionization detection)</td>
			<td>Agilent</td>
			<td></td>
			<td>6890N</td>
		</tr>
		<tr>
			<td>Gastight analytical syringe</td>
			<td>Hamilton</td>
			<td>81220</td>
			<td>1750 TLL</td>
		</tr>
		<tr>
			<td>Gastight analytical syringe needle</td>
			<td>Hamilton</td>
			<td>7729-07</td>
			<td>22 G, metal hub needle, 2 in, point style 5</td>
		</tr>
		<tr>
			<td>Gas-tight vials</td>
			<td>Labco</td>
			<td>938W</td>
			<td>Exetainer vial: 12 mL, round bottom</td>
		</tr>
		<tr>
			<td>Glass culture tubes</td>
			<td>Bellco Glass</td>
			<td>2048-00150</td>
			<td>18 x 150 mm</td>
		</tr>
		<tr>
			<td>LiCl precipitation solution (7.5 M)</td>
			<td>Invitrogen</td>
			<td>AM9480</td>
			<td></td>
		</tr>
		<tr>
			<td>One-way stopcock</td>
			<td>VWR</td>
			<td>MFLX30600-00</td>
			<td>inlet port: female luer, outlet port: male luer lock</td>
		</tr>
		<tr>
			<td>Petri dish, square</td>
			<td>Fisher Scientific</td>
			<td>FB0875711A</td>
			<td>100 x 100 mm</td>
		</tr>
		<tr>
			<td>Phosphate buffer, 0.2 M (pH 6.8)</td>
			<td></td>
			<td>see CAS numbers below</td>
			<td>Dissolve the following in Milli-Q water and autoclave: 12.24 g/L KH2PO4, 26.29 g/L Na2HPO4 &#183; 7H2O</td>
		</tr>
		<tr>
			<td>Pierce BCA Protein Assay Kit&#160;</td>
			<td>Thermo Scientific</td>
			<td>23225</td>
			<td></td>
		</tr>
		<tr>
			<td>PTFE syringe filter tip</td>
			<td>Thermo Scientific</td>
			<td>03-050-469</td>
			<td>hydrophobic, pore size: 0.2 &#181;m, diameter: 4 mm</td>
		</tr>
		<tr>
			<td>Qubit 1x dsDNA High Sensitivity Assay Kit&#160;</td>
			<td>Invitrogen</td>
			<td>Q33230</td>
			<td></td>
		</tr>
		<tr>
			<td>Qubit 4 Fluorometer</td>
			<td>Invitrogen</td>
			<td>Q33238</td>
			<td></td>
		</tr>
		<tr>
			<td>RNA Clean &#38; Concentrator-5</td>
			<td>Zymo Research</td>
			<td>R1013</td>
			<td></td>
		</tr>
		<tr>
			<td>Serum stopper</td>
			<td>Fisher Scientific</td>
			<td>03-340-302</td>
			<td>20 mm</td>
		</tr>
		<tr>
			<td>Syringe</td>
			<td>BD Biosciences</td>
			<td>302995</td>
			<td>Luer-Lock, 10 mL, single use, sterile</td>
		</tr>
		<tr>
			<td>Syringe pump</td>
			<td>New Era Pump Systems Inc.</td>
			<td>1000-US</td>
			<td>NE-1000 one channel programmable</td>
		</tr>
		<tr>
			<td>SYTO9, propidium iodide, microspheres</td>
			<td>Invitrogen</td>
			<td>L34856</td>
			<td>LIVE/DEAD BacLight Bacterial Viability Kit</td>
		</tr>
		<tr>
			<td>Zirconia/silica beads</td>
			<td>BioSpec Products</td>
			<td>11079101z</td>
			<td>0.1 mm diameter</td>
		</tr>
		<tr>
			<td>Chemical reagents</td>
			<td>CAS number</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2&#183;6H2O</td>
			<td>7774-34-7</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CoCl2&#183;6H2O</td>
			<td>7791-13-1</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Concentrated sulfuric acid</td>
			<td>7664-93-9</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CTAB, cetrimonium bromide</td>
			<td>57-09-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>CuCl2&#183;2H2O</td>
			<td>10125-13-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>64-17-5</td>
			<td></td>
			<td>&#160;&#160;</td>
		</tr>
		<tr>
			<td>FeSO4&#183;7H2O</td>
			<td>7782-63-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>H3BO3</td>
			<td>10043-35-3</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Isopropanol</td>
			<td>69-63-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KH2PO4</td>
			<td>7778-77-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>KNO3</td>
			<td>7757-79-1</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>LaCl3</td>
			<td>10099-58-8</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4&#183;7H2O</td>
			<td>10034-99-8</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>MnCl2&#183;4H2O</td>
			<td>13446-34-9</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Na2CO3, sodium carbonate</td>
			<td>497-19-8</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Na2-EDTA</td>
			<td>139-33-3</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Na2HPO4 &#183; 7H2O</td>
			<td>7782-85-6</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Na2MoO&#183;2H2O</td>
			<td>10102-40-6</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NaCl, sodium chloride</td>
			<td>7647-14-5</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>NiCl2&#183;6H2O</td>
			<td>7791-20-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Phenol (90% solution in water)</td>
			<td>108-95-2</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>PVP40, polyvinylpyrrolidone</td>
			<td>9003-39-8</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-HCl</td>
			<td>1185-53-1</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>ZnSO4&#183;7H2O</td>
			<td>7446-20-0</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;-Mercaptoethanol</td>
			<td>60-24-2</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>

agarose-based model ecosystem for cultivating methanotrophs in a methane-oxygen counter gradient

Aerobic methane-oxidizing bacteria, known as methanotrophs, serve important roles in biogeochemical cycling. Methanotrophs occupy a specific environmental niche within methane-oxygen counter gradients found in soils and sediments, which influences their behavior on an individual and community level. However, conventional methods to study the physiology of these greenhouse gas-mitigating microorganisms often use homogeneous planktonic cultures, which do not accurately represent the spatial and chemical gradients found in the environment. This hinders scientists' understanding of how these bacteria behave in situ. Here, a simple, inexpensive model ecosystem called the gradient syringe is described, which uses semi-solid agarose to recreate the steep methane-oxygen counter gradients characteristic of methanotrophs' natural habitats. The gradient syringe allows for the cultivation of methanotrophic strains and the enrichment of mixed methane-oxidizing consortia from environmental samples, revealing phenotypes only visible in this spatially resolved context. This protocol also reports various biochemical assays that have been modified to be compatible with the semi-solid agarose matrix, which may be valuable to researchers culturing microorganisms within other agarose-based systems.

Here, the gradient syringe model ecosystem was used to cultivate a single strain (the methanotroph Methylomonas sp. strain LW13) (Figure 1A)6, but it can also be used to enrich for a methane-oxidizing microbial community by direct soil inoculation (Figure 1B). The presence of a methane-oxygen counter gradient was validated by measuring the concentration of methane and oxygen across cell-free and inoculated syringes (Figure 1C). For LW13-inoculated gradient syringes, a counter gradient formed within one day of flushing the syringe, which steepened over three days of incubation. Over the same time period, a horizontal band formed at the same depth at which both gas substrates reached their lowest concentrations (Figure 1A). The steep gas gradient and depletion of methane and oxygen past the depth of the horizontal band showed that LW13 aerobically metabolized methane and produced a phenotype not observed in homogenous planktonic culture. This phenotype is also produced by other methanotrophic bacteria isolated from the same environmental sample as LW136. Strain-dependent variation in the timing and depth of the horizontal band development among different methanotrophic strains suggested that the horizontal band was affected by the specific behavior of each microbe when cultivated in a spatially resolved context6.
The number of cells in the entire agarose plug was measured using flow cytometry and colony counts (CFU/mL) (Figure 2A). This method was used to compare the cell distribution and survival of wild-type LW13 to a mutant strain of LW13 containing a deletion in the fucose 4-O-acetyltransferase (OAT) gene, which was previously shown to influence horizontal band development6. The &#916;OAT mutant of LW13 had lower overall growth in the gradient syringe over 6 days compared to the wild-type, an effect that was not observed in homogenous planktonic cultures of the same strains6. The mutant strain did not form the same distinct horizontal band as the LW13 wild type when cultured in the gradient syringe (Figure 2B). Cell numbers and horizontal band appearance were restored to levels similar to the wild type upon gene complementation in the mutant strain. These results demonstrate that the gradient syringe can be used to link genes to specific phenotypes only present in the methane-oxygen counter gradient.
A variety of genetic, chemical, and molecular techniques were adapted for use with bacteria grown within a semi-solid agarose matrix. The gradient syringe model ecosystem can be readily used for standard biomolecular quantification assays with the inclusion of uninoculated agarose as the negative control. The concentration of three different biomolecules commonly found in extracellular polymeric substances and biofilms was measured: polysaccharides, protein, and extracellular DNA15 (Figure 3). In LW13-inoculated syringe segments, the horizontal band had significantly more polysaccharides than other segments, with no significant increase in protein or extracellular DNA.
RNA-seq was used to measure transcriptional differences in LW13 growing at different depths of the syringe. Robust RNA extraction was achieved using a CTAB-based extraction buffer followed by conventional phenol: chloroform extraction and precipitation steps. The results from the RNA-seq analysis were later used to identify genes implicated in the production of the horizontal polysaccharide band. These results indicate that the semi-solid agarose essential for the creation of a spatially resolved model ecosystem does not prevent further biochemical analyses that are generally reserved for planktonic and plate-based cultures.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67191/67191fig01.jpg" /> 
Figure 1: The gradient syringe model ecosystem. (A) The gradient syringe inoculated with the methanotroph Methylomonas sp. LW13. A distinct horizontal band (arrowhead) develops within two days of flushing the syringe with 100% methane. (B) Close-up photos of gradient syringes inoculated with soil diluted 10-1 and 10-4 and incubated for two weeks. Gradient syringes containing more dilute soil resulted in spherical colonies throughout the agarose, whereas more concentrated soil inocula resulted in a distinct band. (C) Characterization of the methane-oxygen counter gradient in LW13-inoculated and sterile gradient syringes after three days of incubation. The gray bar indicates the range of depths at which the polysaccharide band was located; data show the mean &#177; SD of three independent experiments with three technical replicates each. Panels (A) and (C) were modified from Beals et al.6. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67191/67191fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67191/67191fig02.jpg" /> 
Figure 2: Quantification of LW13 wild type and mutant after incubation in the gradient syringe. (A) The total number of LW13 cells per mL extruded agarose recovered from gradient syringes on Day 0 and Day 7 measured by flow cytometry. &#916;OAT contains a deletion of the fucose 4-O-acetyltransferase gene, which was found to be highly expressed in cells located at the depth of the polysaccharide band. &#916;OAT+OAT contains the OAT gene inserted at a distal location of the &#916;OAT genome. *, significantly different (two-tailed heteroscedastic t-test, &#945; = 0.05); n.s., not significantly different. Data show the mean &#177; SD of three independent experiments with two technical replicates each. (B) Horizontal band development in gradient syringes inoculated with LW13 wild type, &#916;OAT, or &#916;OAT+OAT after seven days of incubation.&#160;This figure was modified from Beals et al.6.&#160;<a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67191/67191fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67191/67191fig03.jpg" /> 
Figure 3: Quantification of biomolecules at increasing depths of the gradient syringe. Relative polysaccharide content (%), protein concentration (&#181;g/mL), and DNA concentration (ng/&#181;L) in eight sections of LW13-inoculated gradient syringes incubated for seven days (filled circles; open circles show values from sterile gradient syringes). Data show the mean &#177; SD of three independent experiments with three technical replicates each. For relative polysaccharide content, * indicates a significant difference from sterile control at equivalent depth (two-tailed heteroscedastic t-test, &#945; = 0.05). For protein and DNA concentrations, * indicates a significant difference from the section containing the horizontal band (one-way ANOVA with Tukey-Kramer post hoc analysis); n.s., not significant. The figure was&#160;modified from Beals et al.6. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/67191/67191fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Author Spotlight: Designing Simple and Inexpensive Techniques to Grow Methane-Oxidizing Bacteria in the Laboratory

A protocol is described for preparing a simple model ecosystem that recreates the methane-oxygen counter gradient found in the natural habitat of aerobic methane-oxidizing bacteria, enabling the study of their physiology in a spatially resolved context. Modifications to common biochemical assays for use with the agarose-based model ecosystem are also described.

Agarose-Based Model Ecosystem for Cultivating Methanotrophs in a Methane-Oxygen Counter Gradient

Aerobic methane-oxidizing bacteria, known as methanotrophs, serve important roles in biogeochemical cycling. Methanotrophs occupy a specific environmental ...

agarose-based-model-ecosystem-for-cultivating-methanotrophs-methane

Nanyang Technological University

Research

JoVE Journal

Environment

749 Views.  University of Utah. A protocol is described for preparing a simple model ecosystem that recreates the methane-oxygen counter gradient found in the natural habitat of aerobic methane-oxidizing bacteria, enabling the study of their physiology in a spatially resolved context. Modifications to common biochemical assays for use with the agarose-based model ecosystem are also described.

Video: Author Spotlight: Designing Simple and Inexpensive Techniques to Grow Methane-Oxidizing Bacteria in the Laboratory

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail millet. Recently, the 510 Mb genome of foxtail millet, S. italica, has been sequenced 1,2 and a 25x coverage genome sequence of the weedy relative S. viridis is in progress. S. viridis has a number of characteristics that make it a potentially excellent model genetic system including a rapid generation time, small stature, simple growth requirements, prolific seed production 3 and developed systems for both transient and stable transformation 4. However, the genetics of S. viridis is largely unexplored, in part, due to the lack of detailed methods for performing crosses. To date, no standard protocol has been adopted that will permit rapid production of seeds from controlled crosses.
The protocol presented here is optimized for performing genetic crosses in S. viridis, accession A10.1. We have employed a simple heat treatment with warm water for emasculation after pruning the panicle to retain 20-30 florets and labeling of flowers to eliminate seeds resulting from newly developed flowers after emasculation. After testing a series of heat treatments at permissive temperatures and varying the duration of dipping, we have established an optimum temperature and time range of 48 &deg;C for 3-6 min. By using this method, a minimum of 15 crosses can be performed by a single worker per day and an average of 3-5 outcross progeny per panicle can be recovered. Therefore, an average of 45-75 outcross progeny can be produced by one person in a single day. Broad implementation of this technique will facilitate the development of recombinant inbred line populations of S. viridis X S. viridis or S. viridis X S. italica, mapping mutations through bulk segregant analysis and creating higher order mutants for genetic analysis.

Methods for Performing Crosses in Setaria viridis, a New Model System for the Grasses

We have developed a methodology for performing crosses in Setaria viridis (S. viridis). The method involves pruning the panicle prior to a hot water treatment to kill viable pollen. Crosses are performed following a well-controlled growth regime and typically result in the recovery of 1 to 7 cross-pollinated seed/s per panicle.

Setaria viridis is an emerging model system for C4 grasses. It is closely related to the bioenergy feed stock switchgrass and the grain crop foxtail ...

Application of Two-spotted Spider Mite Tetranychus urticae for Plant-pest Interaction Studies

The two-spotted spider mite, Tetranychus urticae, is a ubiquitous polyphagous arthropod herbivore that feeds on a remarkably broad array of species, with more than 150 of economic value. It is a major pest of greenhouse crops, especially in Solanaceae and Cucurbitaceae (e.g., tomatoes, eggplants, peppers, cucumbers, zucchini) and greenhouse ornamentals (e.g., roses, chrysanthemum, carnations), annual field crops (such as maize, cotton, soybean, and sugar beet), and in perennial cultures (alfalfa, strawberries, grapes, citruses, and plums)1,2. In addition to the extreme polyphagy that makes it an important agricultural pest, T. urticae has a tendency to develop resistance to a wide array of insecticides and acaricides that are used for its control3-7.
T. urticae is an excellent experimental organism, as it has a rapid life cycle (7 days at 27 &#176;C) and can be easily maintained at high density in the laboratory. Methods to assay gene expression (including in situ hybridization and antibody staining) and to inactivate expression of spider mite endogenous genes using RNA interference have been developed8-10. Recently, the whole genome sequence of T. urticae has been reported, creating an opportunity to develop this pest herbivore as a model organism with equivalent genomic resources that already exist in some of its host plants (Arabidopsis thaliana and the&#160;tomato Solanum lycopersicum)11. Together, these&#160;model organisms could provide insights into molecular bases of plant-pest interactions.
Here, an efficient method for quick and easy collection of a large number of adult female mites, their application on an experimental plant host, and the assessment of the plant damage due to spider mite feeding are described. The presented protocol enables fast and efficient collection of hundreds of individuals at any developmental stage (eggs, larvae, nymphs, adult males, and females) that can be used for subsequent experimental application.

Application of Two-spotted Spider Mite Tetranychus urticae for Plant-pest Interaction Studies

Protocols for efficient preparation of homogenous samples of spider mites, infestation of experimental plants, and assessment of plant damage, as required for studies of plant-pest interaction were developed.

The two-spotted spider mite, Tetranychus urticae, is a ubiquitous polyphagous arthropod herbivore that feeds on a remarkably broad array of species, with ...

Removal of Trace Elements by Cupric Oxide Nanoparticles from Uranium In Situ Recovery Bleed Water and Its Effect on Cell Viability

In situ recovery (ISR) is the predominant method of uranium extraction in the United States. During ISR, uranium is leached from an ore body and extracted through ion exchange. The resultant production bleed water (PBW) contains contaminants such as arsenic and other heavy metals. Samples of PBW from an active ISR uranium facility were treated with cupric oxide nanoparticles (CuO-NPs). CuO-NP treatment of PBW reduced priority contaminants, including arsenic, selenium, uranium, and vanadium. Untreated and CuO-NP treated PBW was used as the liquid component of the cell growth media and changes in viability were determined by the MTT (3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay in human embryonic kidney (HEK 293) and human hepatocellular carcinoma (Hep G2) cells. CuO-NP treatment was associated with improved HEK and HEP cell viability. Limitations of this method include dilution of the PBW by growth media components and during osmolality adjustment as well as necessary pH adjustment. This method is limited in its wider context due to dilution effects and changes in the pH of the PBW which is traditionally slightly acidic&#160;however; this method could have a broader use assessing CuO-NP treatment in more neutral waters.

Removal of Trace Elements by Cupric Oxide Nanoparticles from Uranium In Situ Recovery Bleed Water and Its Effect on Cell Viability

Production bleed water (PBW) was treated with cupric oxide nanoparticles (CuO-NPs) and cellular toxicity was assessed in cultured human cells. The goal of this protocol was to integrate the native environmental sample into a cell culture format assessing the changes in toxicity due to CuO-NP treatment.

In situ recovery (ISR) is the predominant method of uranium extraction in the United States. During ISR, uranium is leached from an ore body and extracted ...

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

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

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

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

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

Screening for Endocrine Activity in Water Using Commercially-available In Vitro Transactivation Bioassays

In vitro transactivation bioassays have shown promise as water quality monitoring tools, however their adoption and widespread application has been hindered partly due to a lack of standardized methods and availability of robust, user-friendly technology. In this study, commercially available, division-arrested cell lines were employed to quantitatively screen for endocrine activity of chemicals present in water samples of interest to environmental quality professionals. A single, standardized protocol that included comprehensive quality assurance/quality control (QA/QC) checks was developed for Estrogen and Glucocorticoid Receptor activity (ER and GR, respectively) using a cell-based Fluorescence Resonance Energy Transfer (FRET) assay. Samples of treated municipal wastewater effluent and surface water from freshwater systems in California (USA), were extracted using solid phase extraction and analyzed for endocrine activity using the standardized protocol. Background and dose-response for endpoint-specific reference chemicals met QA/QC guidelines deemed necessary for reliable measurement. The bioassay screening response for surface water samples was largely not detectable. In contrast, effluent samples from secondary treatment plants had the highest measurable activity, with estimated bioassay equivalent concentrations (BEQs) up to 392 ng dexamethasone/L for GR and 17 ng 17&#946;-estradiol/L for ER. The bioassay response for a tertiary effluent sample was lower than that measured for secondary effluents, indicating a lower residual of endocrine active chemicals after advanced treatment. This protocol showed that in vitro transactivation bioassays that utilize commercially available, division-arrested cell &#34;kits&#34;, can be adapted to screen for endocrine activity in water.

Screening for Endocrine Activity in Water Using Commercially-available In Vitro Transactivation Bioassays

A protocol to screen for endocrine activity in organic extracts of water samples, including treated wastewater effluent and surface (receiving) water, was adapted using commercially available division-arrested ("freeze and thaw") in vitro transactivation bioassays.

In vitro transactivation bioassays have shown promise as water quality monitoring tools, however their adoption and widespread application has been ...

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and rates of mortality in populations, which has a variety of applications in ecology, including agricultural ecosystems. Horizontal, or cohort-based, life tables provide for the most direct and accurate method of quantifying vital population rates because they follow a group of individuals in a population from birth to death. Here, protocols are presented for conducting and analyzing cohort-based life tables in the field that takes advantage of the sessile nature of the immature life stages of a global insect pest, Bemisia tabaci. Individual insects are located on the underside of cotton leaves and are marked by drawing a small circle around the insect with a non-toxic pen. This insect can then be observed repeatedly over time with the aid of hand lenses to measure development from one stage to the next and to identify stage-specific causes of death associated with natural and introduced mortality forces. Analyses explain how to correctly measure multiple mortality forces that act contemporaneously within each stage and how to use such data to provide meaningful population dynamic metrics. The method does not directly account for adult survival and reproduction, which limits inference to dynamics of immature stages. An example is presented that focused on measuring the impact of bottom-up (plant quality) and top-down (natural enemies) effects on the mortality dynamics of B. tabaci in the cotton system.

Methodology for Developing Life Tables for Sessile Insects in the Field Using the Whitefly, Bemisia tabaci, in Cotton As a Model System

Life tables allow quantification of the sources and rates of mortality in insect populations and contribute to understanding, predicting and manipulating population dynamics in agroecosystems. Methods for conducting and analyzing cohort-based life tables in the field for an insect with sessile immature life stages are presented.

Life tables provide a means of measuring the schedules of birth and death from populations over time. They also can be used to quantify the sources and ...

Continuous Hydrologic and Water Quality Monitoring of Vernal Ponds

Vernal ponds, also referred to as vernal pools, provide critical ecosystem services and habitat for a variety of threatened and endangered species. However, they are vulnerable parts of the landscapes that are often poorly understood and understudied. Land use and management practices, as well as climate change are thought to be a contribution to the global amphibian decline. However, more research is needed to understand the extent of these impacts. Here, we present methodology for characterizing a vernal pond's morphology and detail a monitoring station that can be used to collect water quantity and quality data over the duration of a vernal pond's hydroperiod. We provide methodology for how to conduct field surveys to characterize the morphology and develop stage-storage curves for a vernal pond. Additionally, we provide methodology for monitoring the water level, temperature, pH, oxidation-reduction potential, dissolved oxygen, and electrical conductivity of water in a vernal pond, as well as monitoring rainfall data. This information can be used to better quantify the ecosystem services that vernal ponds provide and the impacts of anthropogenic activities on their ability to provide these services.

Understanding the ecosystem services and processes provided by vernal ponds and the impacts of anthropogenic activities on their ability to provide these services requires intensive hydrologic monitoring. This sampling protocol using in-situ monitoring equipment was developed to understand the impact of anthropogenic activities on water levels and quality.

Vernal ponds, also referred to as vernal pools, provide critical ecosystem services and habitat for a variety of threatened and endangered species. ...

Ecosystem Fabrication (EcoFAB) Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A better mechanistic understanding of these complex plant-microbe interactions will be crucial to improving plant production as well as performing basic ecological studies investigating plant-soil-microbe interactions. Here, a detailed description for ecosystem fabrication is presented, using widely available 3D printing technologies, to create controlled laboratory habitats (EcoFABs) for mechanistic studies of plant-microbe interactions within specific environmental conditions. Two sizes of EcoFABs are described that are suited for the investigation of microbial interactions with various plant species, including Arabidopsis thaliana, Brachypodium distachyon, and Panicum virgatum. These flow-through devices allow for controlled manipulation and sampling of root microbiomes, root chemistry as well as imaging of root morphology and microbial localization. This protocol includes the details for maintaining sterile conditions inside EcoFABs and mounting independent LED light systems onto EcoFABs. Detailed methods for addition of different forms of media, including soils, sand, and liquid growth media coupled to the characterization of these systems using imaging and metabolomics are described. Together, these systems enable dynamic and detailed investigation of plant and plant-microbial consortia including the manipulation of microbiome composition (including mutants), the monitoring of plant growth, root morphology, exudate composition, and microbial localization under controlled environmental conditions. We anticipate that these detailed protocols will serve as an important starting point for other researchers, ideally helping create standardized experimental systems for investigating plant-microbe interactions.

Ecosystem Fabrication EcoFAB Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

This article describes detailed protocols for ecosystem fabrication of devices (EcoFABs) that enable the studies of plants and plant-microbe interactions in highly controlled laboratory conditions.

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A ...

Wastewater Irrigation Impacts on Soil Hydraulic Conductivity: Coupled Field Sampling and Laboratory Determination of Saturated Hydraulic Conductivity

Since the early 1960s, an alternative wastewater discharge practice at The Pennsylvania State University has been researched and its impacts monitored. Rather than discharging treated wastewater to a stream, and thereby directly impacting the stream quality, the effluent is applied to forested and cropped land managed by the University. Concerns related to reductions in soil hydraulic conductivity occur when considering wastewater reuse. The methodology described in this manuscript, matching soil sample size with the size of the laboratory-based hydraulic conductivity measurement apparatus, provides the benefits of a relatively rapid collection of samples with the benefits of controlled laboratory boundary conditions. The results suggest that there may have been some impact of wastewater reuse on the soil&#39;s ability to transmit water at deeper depths in the depressional areas of the site. Most of the reductions in the soil hydraulic conductivity in the depressions appear to be related to the depth from which the sample was collected, and by inference, associated with the soil structural and textural differences.

Here we present a methodology which matches a soil sample size and a hydraulic conductivity measurement device to prevent the so-called wall flow along the inside of the soil container from being erroneously included in water flow measurements. Its use is demonstrated with samples collected from a wastewater irrigation site.

Since the early 1960s, an alternative wastewater discharge practice at The Pennsylvania State University has been researched and its impacts monitored. ...

The Hawaii Protocol for Scientific Monitoring of Coffee Berry Borer: a Model for Coffee Agroecosystems Worldwide

Coffee berry borer (CBB) is the most devastating insect pest for coffee crops worldwide. We developed a scientific monitoring protocol that is aimed at capturing and quantifying the dynamics and impact of this invasive insect pest as well as the development of its host plant across a heterogeneous landscape. The cornerstone of this comprehensive monitoring system is timely georeferenced data collection on CBB movement, coffee berry infestation, mortality by the fungus Beauveria bassiana, and coffee plant phenology via a mobile electronic data recording application. This electronic data collection system allows field records to be georeferenced through built-in global positioning systems, and is backed by a network of weather stations and records of farm management practices. Comprehensive monitoring of CBB and host plant dynamics is an essential part of an area-wide project in Hawaii to aggregate landscape-level data for research to improve management practices. Coffee agroecosystems in other parts of the world that experience highly variable environmental and socioeconomic factors will also benefit from implementing this protocol, in that it will drive the development of customized integrated pest management (IPM) to manage CBB populations.

Comprehensive monitoring of coffee berry borer and host plant dynamics is essential for aggregating landscape-level data to improve management of this invasive pest. Here, we present a protocol for scientific monitoring of coffee berry borer movement, infestation, mortality, coffee plant phenology, weather, and farm management via a mobile electronic data recording application.

Coffee berry borer (CBB) is the most devastating insect pest for coffee crops worldwide. We developed a scientific monitoring protocol that is aimed at ...