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><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&amp;nbsp;</td><td></td><td>see CAS numbers below</td><td>Dissolve the following in Mili-Q water and autoclave:&amp;nbsp; 0.2 g/L MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O, 0.2 g/L CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O, 1 g/L KNO&lt;sub&gt;3&lt;/sub&gt;, and 30 &amp;mu;M LaCl&lt;sub&gt;3&lt;/sub&gt;. 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 Na&lt;sub&gt;2&lt;/sub&gt;-EDTA, 2.0 g/L FeSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O, 0.8 g/L ZnSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O, 0.03 g/L MnCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;4H&lt;sub&gt;2&lt;/sub&gt;O, 0.03 g/L H&lt;sub&gt;3&lt;/sub&gt;BO&lt;sub&gt;3&lt;/sub&gt;, 0.2 g/L CoCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O, 0.6 g/L CuCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O, 0.02 g/L NiCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O, and 0.05 g/L Na&lt;sub&gt;2&lt;/sub&gt;MoO&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O.</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>&amp;nbsp;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 O&lt;sub&gt;2&lt;/sub&gt; 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 KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;, 26.29 g/L Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; &amp;middot; 7H&lt;sub&gt;2&lt;/sub&gt;O</td></tr><tr><td>Pierce BCA Protein Assay Kit&amp;nbsp;</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 &amp;micro;m, diameter: 4 mm</td></tr><tr><td>Qubit 1x dsDNA High Sensitivity Assay Kit&amp;nbsp;</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 &amp;amp; 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>&lt;strong&gt;Chemical reagents&lt;/strong&gt;</td><td>&lt;strong&gt;CAS number&lt;/strong&gt;</td><td></td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>7774-34-7</td><td></td><td></td></tr><tr><td>CoCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</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>CuCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>10125-13-0</td><td></td><td></td></tr><tr><td>Ethanol</td><td>64-17-5</td><td></td><td>&amp;nbsp;&amp;nbsp;</td></tr><tr><td>FeSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>7782-63-0</td><td></td><td></td></tr><tr><td>H&lt;sub&gt;3&lt;/sub&gt;BO&lt;sub&gt;3&lt;/sub&gt;</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>KH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;</td><td>7778-77-0</td><td></td><td></td></tr><tr><td>KNO&lt;sub&gt;3&lt;/sub&gt;</td><td>7757-79-1</td><td></td><td></td></tr><tr><td>LaCl&lt;sub&gt;3&lt;/sub&gt;</td><td>10099-58-8</td><td></td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>10034-99-8</td><td></td><td></td></tr><tr><td>MnCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;4H&lt;sub&gt;2&lt;/sub&gt;O</td><td>13446-34-9</td><td></td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;CO&lt;sub&gt;3&lt;/sub&gt;, sodium carbonate</td><td>497-19-8</td><td></td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;-EDTA</td><td>139-33-3</td><td></td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt; &amp;middot; 7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>7782-85-6</td><td></td><td></td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;MoO&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</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>NiCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</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>ZnSO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>7446-20-0</td><td></td><td></td></tr><tr><td>&amp;beta;-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/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/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/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

Research

JoVE Journal

Environment

781 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

Development of Sulfidogenic Sludge from Marine Sediments and Trichloroethylene Reduction in an Upflow Anaerobic Sludge Blanket Reactor

The importance of microbial sulfate reduction relies on the various applications that it offers in environmental biotechnology. Engineered sulfate reduction is used in industrial wastewater treatment to remove large concentrations of sulfate along with the chemical oxygen demand (COD) and heavy metals. The most common approach to the process is with anaerobic bioreactors in which sulfidogenic sludge is obtained through adaptation of predominantly methanogenic granular sludge to sulfidogenesis. This process may take a long time and does not always eliminate the competition for substrate due to the presence of methanogens in the sludge. In this work, we propose a novel approach to obtain sulfidogenic sludge in which hydrothermal vents sediments are the original source of microorganisms. The microbial community developed in the presence of sulfate and volatile fatty acids is wide enough to sustain sulfate reduction over a long period of time without exhibiting inhibition due to sulfide. 
This protocol describes the procedure to generate the sludge from the sediments in an upflow anaerobic sludge blanket (UASB) type of reactor. Furthermore, the protocol presents the procedure to demonstrate the capability of the sludge to remove by reductive dechlorination a model of a highly toxic organic pollutant such as trichloroethylene (TCE). The protocol is divided in three stages: (1) the formation of the sludge and the determination of its sulfate reducing activity in the UASB, (2) the experiment to remove the TCE by the sludge, and (3) the identification of microorganisms in the sludge after the TCE reduction. Although in this case the sediments were taken from a site located in Mexico, the generation of a sulfidogenic sludge by using this procedure may work if a different source of sediments is taken since marine sediments are a natural pool of microorganisms that may be enriched in sulfate reducing bacteria.

Microbial sulfate reduction is a process of great importance in environmental biotechnology. The success of the sulfidogenic reactors depends among other factors on the microbial composition of the sludge. Here, we present a protocol to develop sulfidogenic sludge from hydrothermal vents sediments in a UASB reactor for reductive dechlorination purposes.

The importance of microbial sulfate reduction relies on the various applications that it offers in environmental biotechnology. Engineered sulfate ...

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in nutrient and gas cycles. After cores with intact sediment-water interfaces are collected, they are submerged in incubation tanks and kept under aerobic conditions at in situ temperatures. To initiate a time course of overlying water chemistry, cores are sealed without bubbles using a top cap with a suspended stirrer. Time courses of 4-7 sample points are used to determine the rate of sediment water exchange. Artificial illumination simulates day-time conditions for shallow photosynthetic sediments, and in conjunction with dark incubations can provide net exchanges on a daily basis. The net measurement of N2 is made possible by sampling a time course of dissolved gas concentrations, with high precision mass spectrometric analysis of N2:Ar ratios providing a means to measure N2 concentrations. We have successfully applied this approach to lakes, reservoirs, estuaries, wetlands and storm water ponds, and with care, this approach provides valuable information on biogeochemical balances in aquatic ecosystems.

The Benthic Exchange of O2, N2 and Dissolved Nutrients Using Small Core Incubations

Here, we present small core incubations for the measurement of sediment-water gas and solute exchange. These will provide reliable measurements of sediment-water exchange that assess the role of sediment in influencing biological and biogeochemical processes in aquatic ecosystems.

The measurement of sediment-water exchange of gases and solutes in aquatic sediments provides data valuable for understanding the role of sediments in ...

Laboratory Simulation of an Iron(II)-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling from hydrothermal sources in the Precambrian ocean was oxidized by molecular oxygen [O2] produced by cyanobacteria. The oldest BIFs, deposited prior to the Great Oxidation Event (GOE) at about 2.4 billion years (Gy) ago, could have formed by direct oxidation of Fe(II) by anoxygenic photoferrotrophs under anoxic conditions. As a method for testing the geochemical and mineralogical patterns that develop under different biological scenarios, we designed a 40 cm long vertical flow-through column to simulate an anoxic Fe(II)-rich marine upwelling system representative of an ancient ocean on a lab scale. The cylinder was packed with a porous glass bead matrix to stabilize the geochemical gradients, and liquid samples for iron quantification could be taken throughout the water column. Dissolved oxygen was detected non-invasively via optodes from the outside. Results from biotic experiments that involved upwelling fluxes of Fe(II) from the bottom, a distinct light gradient from top, and cyanobacteria present in the water column, show clear evidence for the formation of Fe(III) mineral precipitates and development of a chemocline between Fe(II) and O2. This column allows us to test hypotheses for the formation of the BIFs by culturing cyanobacteria (and in the future photoferrotrophs) under simulated marine Precambrian conditions. Furthermore we hypothesize that our column concept allows for the simulation of various chemical and physical environments &#8212; including shallow marine or lacustrine sediments.

Laboratory Simulation of an IronII-rich Precambrian Marine Upwelling System to Explore the Growth of Photosynthetic Bacteria

We simulated a Precambrian ferruginous marine upwelling system in a lab-scale vertical flow-through column. The goal was to understand how geochemical profiles of O2 and Fe(II) evolve as cyanobacteria produce O2. The results show the establishment of a chemocline due to Fe(II) oxidation by photosynthetically produced O2.

A conventional concept for the deposition of some Precambrian Banded Iron Formations (BIF) proceeds on the assumption that ferrous iron [Fe(II)] upwelling ...

Continuously-stirred Anaerobic Digester to Convert Organic Wastes into Biogas: System Setup and Basic Operation

Anaerobic digestion (AD) is a bioprocess that is commonly used to convert complex organic wastes into a useful biogas with methane as the energy carrier 1-3. Increasingly, AD is being used in industrial, agricultural, and municipal waste(water) treatment applications 4,5. The use of AD technology allows plant operators to reduce waste disposal costs and offset energy utility expenses. In addition to treating organic wastes, energy crops are being converted into the energy carrier methane 6,7. As the application of AD technology broadens for the treatment of new substrates and co-substrate mixtures 8, so does the demand for a reliable testing methodology at the pilot- and laboratory-scale.
Anaerobic digestion systems have a variety of configurations, including the continuously stirred tank reactor (CSTR), plug flow (PF), and anaerobic sequencing batch reactor (ASBR) configurations 9. The CSTR is frequently used in research due to its simplicity in design and operation, but also for its advantages in experimentation. Compared to other configurations, the CSTR provides greater uniformity of system parameters, such as temperature, mixing, chemical concentration, and substrate concentration. Ultimately, when designing a full-scale reactor, the optimum reactor configuration will depend on the character of a given substrate among many other nontechnical considerations. However, all configurations share fundamental design features and operating parameters that render the CSTR appropriate for most preliminary assessments. If researchers and engineers use an influent stream with relatively high concentrations of solids, then lab-scale bioreactor configurations cannot be fed continuously due to plugging problems of lab-scale pumps with solids or settling of solids in tubing. For that scenario with continuous mixing requirements, lab-scale bioreactors are fed periodically and we refer to such configurations as continuously stirred anaerobic digesters (CSADs).
This article presents a general methodology for constructing, inoculating, operating, and monitoring a CSAD system for the purpose of testing the suitability of a given organic substrate for long-term anaerobic digestion. The construction section of this article will cover building the lab-scale reactor system. The inoculation section will explain how to create an anaerobic environment suitable for seeding with an active methanogenic inoculum. The operating section will cover operation, maintenance, and troubleshooting. The monitoring section will introduce testing protocols using standard analyses. The use of these measures is necessary for reliable experimental assessments of substrate suitability for AD. This protocol should provide greater protection against a common mistake made in AD studies, which is to conclude that reactor failure was caused by the substrate in use, when really it was improper user operation 10.

Laboratory-scale anaerobic digesters allow scientists to research new ways of optimizing existing applications of anaerobic biotechnology and to evaluate the methane producing potential of various organic wastes. This article introduces a generalized model for the construction, inoculation, operation, and monitoring of a laboratory-scale continuously stirred anaerobic digester.

Anaerobic digestion (AD) is a bioprocess that is commonly used to convert complex organic wastes into a useful biogas with methane as the energy carrier ...

Bioengineering

Self-standing Electrochemical Set-up to Enrich Anode-respiring Bacteria On-site

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be critical for microbial energy production and persistence in many subsurface environments, especially those lacking soluble terminal electron acceptors. While EET-capable microbes have been successfully isolated from various environments, the diversity of bacteria capable of EET is still poorly understood, especially in difficult-to-sample, low energy or extreme environments, such as many subsurface ecosystems. Here, we describe an on-site electrochemical system to enrich EET-capable bacteria using an anode as a respiratory terminal electron acceptor. This anode is connected&#160;to a cathode capable of&#160;catalyzing abiotic oxygen reduction. Comparing this approach with electrocultivation methods that use a potentiostat for poising the electrode potential, the two-electrode system does not require an external power source. We present an example of our on-site enrichment utilized in an alkaline pond at the Cedars, a terrestrial serpentinization site in Northern California. Prior attempts to cultivate mineral reducing bacteria were unsuccessful, which is likely due to the low-biomass nature of this site and/or the low relative abundance of metal reducing microbes. Prior to implementing our two-electrode enrichment, we measured the vertical profile of dissolved oxygen concentration. This allowed us to place the carbon felt anode and platinum-electroplated carbon felt cathode at depths that would support anaerobic and aerobic processes, respectively. Following on-site incubation, we further enriched the anodic electrode in the laboratory and confirmed&#160;a distinct microbial community compared to the surface-attached or biofilm communities normally observed at the Cedars. This enrichment subsequently led to the isolation of the first electrogenic microbe from the Cedars. This method of on-site microbial enrichment has the potential to greatly enhance the isolation of EET-capable bacteria from low biomass or difficult to sample habitats.

On-site microbial enrichment or in situ cultivation techniques can facilitate the isolation of difficult-to-culture microbial taxa, especially from low-biomass or geochemically extreme environments. Here, we describe an electrochemical set-up without using an external power source to enrich microbial strains that are capable of extracellular electron transport (EET).

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be ...

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

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

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

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

Medium Preparation for the Cultivation of Microorganisms under Strictly Anaerobic/Anoxic Conditions

In contrast to aerobic organisms, strictly anaerobic microorganisms require the absence of oxygen and usually a low redox potential to initiate growth. As oxygen is ubiquitous in air, retaining O2-free conditions during all steps of cultivation is challenging but a prerequisite for anaerobic culturing. The protocol presented here demonstrates the successful cultivation of an anaerobic mixed culture derived from a biogas plant using a simple and inexpensive method. A precise description of the entire anoxic culturing process is given including media preparation, filling of cultivation flasks, supplementation with redox indicator and reducing agents to provide low redox potentials as well as exchanging the headspace to keep media free from oxygen. Furthermore, a detailed overview of aseptically inoculating gas tight serum flasks (by using sterile syringes and needles) and suitable incubation conditions is provided. The present protocol further deals with gas and liquid sampling for subsequent analyses regarding gas composition and volatile fatty acid concentrations using gas chromatography (GC) and high performance liquid chromatography (HPLC), respectively, and the calculation of biogas and methane yield considering the ideal gas law.

As obligate anaerobic organisms are unable to grow upon oxygen exposure, the use of anaerobic culturing techniques is indispensable. Here, we demonstrate a simple and effective method to cultivate a mixed culture derived from a biogas plant from media preparation to gas and volatile fatty acid quantification.

In contrast to aerobic organisms, strictly anaerobic microorganisms require the absence of oxygen and usually a low redox potential to initiate growth. As ...

Biochemistry

In Situ Simulated Anaerobic Method for Culturing Environmental Microbes

A Set of In Situ Informed Simulated Medium Formats for Culturing Environmentally Acquired Anaerobic Microorganisms

Culture-dependent research of anaerobic microorganisms rests upon methodological competence. These methods must create and maintain suitable growth conditions (e.g., pH and carbon sources) for anaerobic microorganisms while also allowing samples to be extracted without compromising the artificial environment. To this end, methods that are informed by and simulate an in situ environment can be of great aid in culturing microorganisms from that environment. Here, we outline an in situ informed and simulated anaerobic method for culturing terrestrial surface and subsurface microorganisms, emphasizing anaerobic sample collection with minimal perturbation. This protocol details the production of a customizable anaerobic liquid medium, and the environmental&#160;acquisition and in vitro growth of anaerobic microorganisms. The protocol also covers critical components of an anaerobic bioreactor used for environmental simulations of sediment and anaerobic liquid media for environmentally acquired cultures. We have included preliminary Next Generation Sequencing data from a maintained microbiome over the lifespan of a bioreactor where the active culture dynamically adjusted in response to an experimental carbon source.

Author Spotlight: Unraveling the Mysteries of Terrestrial Anaerobic Microorganisms in Uncharted Environments by In Situ Culturing

The focus of this paper is to detail best practices for making media for fastidious anaerobic microorganisms acquired from an environment. These methods help manage anaerobic cultures and can be applied to support the growth of elusive uncultured microorganisms, the "microbial dark matter."

Culture-dependent research of anaerobic microorganisms rests upon methodological competence. These methods must create and maintain suitable growth ...

Microalgal Cultivation for Biomethane Obtention from Swine Waste Biogas

Biogas Purification through the use of a Microalgae-Bacterial System in Semi-Industrial High Rate Algal Ponds

In recent years, a number of technologies have emerged to purify biogas into biomethane. This purification entails a reduction in the concentration of polluting gases such as carbon dioxide and hydrogen sulfide to increase the content of methane. In this study, we used a microalgal cultivation technology to treat and purify biogas produced from organic waste from the swine industry to obtain ready-to-use biomethane. For cultivation and purification, two 22.2 m3 open-pond photobioreactors coupled with an absorption-desorption column system were set up in San Juan de los Lagos, Mexico. Several recirculation liquid/biogas ratios (L/G) were tested to obtain the highest removal efficiencies; other parameters, such as pH, dissolved oxygen (DO), temperature, and biomass growth, were measured. The most efficient L/Gs were 1.6 and 2.5, resulting in a treated biogas effluent with a composition of 6.8%vol and 6.6%vol in CO2, respectively, and removal efficiencies for H2S up to 98.9%, as well as maintaining O2 contamination values of less than 2%vol. We found that pH greatly determines CO2 removal, more so than L/G, during cultivation because of its participation in the photosynthetic process of microalgae and its ability to vary pH when solubilized due to its acidic nature. DO, and temperature oscillated as expected from the light-dark natural cycles of photosynthesis and the time of day, respectively. Biomass growth varied with CO2 and nutrient feeding as well as reactor harvesting; however, the trend remained primed for growth.

Author Spotlight: Scaling Microalgal Biotechnology for Enhanced Biomethane Production 

Air pollution impacts the quality of life of all organisms. Here, we describe the use of microalgae biotechnology for the treatment of biogas (simultaneous removal of carbon dioxide and hydrogen sulfide) and the production of biomethane through semi-industrial open high-rate algal ponds and subsequent analysis of treatment efficiency, pH, dissolved oxygen, and microalgae growth.

In recent years, a number of technologies have emerged to purify biogas into biomethane. This purification entails a reduction in the concentration of ...

Visualizing Methane-Cycling Microbial Dynamics in Coastal Wetlands

Coastal wetlands are the largest biotic source of methane, where methanogens convert organic matter into methane and methanotrophs oxidize methane, thus playing a critical role in regulating the methane cycle. The wetlands in South Texas, which are subject to frequent weather events, fluctuating salinity levels, and anthropogenic activities due to climate change, influence methane cycling. Despite the ecological importance of these processes, methane cycling in South Texas coastal wetlands remains insufficiently explored. To address this gap, we developed and optimized a method for detecting genes related to methanogens and methanotrophs, including mcrA as a biomarker for methanogens and pmoA1, pmoA2, and mmoX as biomarkers for methanotrophs. Additionally, this study aimed to visualize the spatial and temporal distribution patterns of methanogen and methanotroph abundance utilizing the geographic information system (GIS) software ArcGIS Pro. The integration of these molecular techniques with advanced geospatial visualization provided critical insights into the spatial and temporal distribution of methanogen and methanotroph communities across South Texas wetlands. Thus, the methodology established in this study offers a robust framework for mapping microbial dynamics in wetlands, enhancing our understanding of methane cycling under varying environmental conditions, and supporting broader ecological and environmental change studies.

The protocol detects key methane-cycling genes in South Texas coastal wetlands and visualizes their spatial distribution to enhance understanding of methane regulation and its environmental impacts in these dynamic ecosystems.

Coastal wetlands are the largest biotic source of methane, where methanogens convert organic matter into methane and methanotrophs oxidize methane, thus ...