Funding for aspects of this work was provided by various sources including the United States Department of Agriculture Specific Cooperative Agreement 58-6408-1-595 and the National Science Foundation (award 1049911).

Colorimetric Analysis of Extracellular Enzyme Activity in Soils and Sediments
1. Preparation of Substrate and Buffer Solutions for Colorimetric Analyses of Enzyme Activity
<ol>
	<li>Prepare 50 mM acetate buffer (pH 5.0-5.5) by mixing 50 ml 0.1 M acetic acid (2.87 ml glacial acetic acid in 500 ml water), 150 ml 0.1 M sodium acetate, and 200 ml distilled H2O. Adjust pH to 5.0-5.5 with 0.1 M acetic acid if necessary.</li>
	<li>Prepare a solution of 1 M sodium hydroxide (NaOH) in distilled H2O.</li>
	<li>Prepare pNP-linked substrate solutions in 50 mM acetate buffer. To assay phosphatase prepare 5 mM pNP-phosphate in 50 mM acetate buffer; to assay &beta;-glucosidase prepare 5 mM pNP-&beta;-glucopyranoside; to assay NAGase prepare 2 mM pNP-&beta;-N-acetylglucosaminide. Prepare all substrate solutions in sterile 15 ml or 50 ml centrifuge tubes. Solutions can be stored at 4 &deg;C for 2-3 weeks.</li>
</ol>
2. Determination of a Standard to Convert Absorbance to pNP Concentration
<ol>
	<li>Prepare standard solutions of p-nitrophenol in 50 mM acetate buffer. Concentrations should range from 0.025-1 mM.</li>
	<li>Transfer three replicates of 100 &mu;l of each concentration to a clear 96-well microplate. Add 10 &mu;l 1 M NaOH and 190 &mu;l distilled H2O to each well.</li>
	<li>Record absorbance at 410 nm using a microplate reader.</li>
	<li>Multiply concentrations in the standard curve by 0.3 to get &mu;moles pNP per 300 &mu;l reaction volume. Plot a curve of absorbance vs. &mu;mole of pNP. The slope of the curve serves as the conversion factor (C) that will relate absorbance to &mu;mole of pNP in each enzyme reaction.</li>
</ol>
3. Conducting the Enzyme Assay
<ol>
	<li>For soil, prepare a slurry of each sample to be assayed in a sterile 15 ml centrifuge tube at a concentration of approximately 1 g/ml-1 using 50 mM acetate buffer. For sediments, add enough acetate buffer to make the slurry easily pipettable. The exact volume of slurry needed will vary according to the number of enzymes assayed, but a minimum of 5 ml is recommended. Vortex each slurry until all clumps of soil or sediment have dispersed and note the final volume.</li>
	<li>Re-vortex each slurry and immediately pipette 150 &mu;l into each of six wells on a 96-well deepwell block (Figure 1). It is important to vortex thoroughly and frequently in order to keep soil particles in suspension. Leave at least two wells per block empty to serve as a substrate control. Note: clip the end of pipette tips with scissors prior to pipetting as soil slurries tend to clog tips. Prepare one 96-well block for each enzyme to be assayed.</li>
	<li>Pour approximately 5 ml of acetate buffer into a pipette reservoir and use an 8-channel pipettor to add 150 &mu;l of the buffer to the last two wells of each sample (these will be sample buffer controls) and the two substrate control wells (Figure 1)</li>
	<li>Pour approximately 10 ml of the appropriate pNP-substrate solution into a pipette reservoir and use an 8-channel pipettor to add 150 &mu;l of the substrate solution to the first four wells of each sample and the two substrate control wells (Figure 1). Note the time as the reaction begins as soon as the substrate solution is added.</li>
	<li>Incubate plates at RT (22 &deg;C) for 0.5-4 hr. Exact incubation time will vary depending on activity level in samples and the enzyme to be assayed. For most soils and sediments, phosphatase and &beta;-glucosidase require incubation times of 0.5-1.5 hr, whereas NAGase and other enzymes require incubation times of &gt;2 hr.</li>
	<li>While assays are incubating prepare clear 96-well microplates to read absorbance. Prepare one microplate for each deepwell block (i.e. for each enzyme assayed). Pipette 10 &mu;l 1 M NaOH and 190 &mu;l of distilled water into each well of the microplate. Note: NaOH slows the enzymatic reaction and raises the pH which enhances the color of the pNP released during the reaction.</li>
	<li>After incubation, centrifuge the 96-well blocks at 2,000-5,000 x g for 5 min to pellet soil particles.</li>
	<li>Use a multichannel pipette to withdraw 100 &mu;l from each well, being careful to avoid the pellet, and transfer it to the corresponding well on prepared clear 96-well microplate.</li>
	<li>Turn on the microplate reader and set up any necessary software. Record absorbance at 410 nm. If the absorbance of a particular well is above the linear detection limit of the plate reader, dilute that well 1:1 with water and re-measure. If absorbance is still too high, the assay should be repeated with a shorter incubation time.</li>
</ol>
4. Determination of Dry Mass of Samples
<ol>
	<li>Pipette 1 ml of each sample slurry into a preweighed aluminum pan.</li>
	<li>Dry in a 75 &deg;C drying oven for 48 hr and weigh. Subtract the weight of the pan from this value to obtain the dry mass of soil or sediment in 1 ml of the slurry. Multiply by a factor of 0.15 to determine the dry mass of sample in the 150 &mu;l added to each well in the enzyme assay.</li>
</ol>
5. Calculation of Enzyme Activity per Dry Mass of Soil or Sediment
<ol>
	<li>Calculate final absorbance of each sample by subtracting the sample control absorbance from the sample assay absorbance. If substrate controls have high absorbance (roughly &gt;0.060) then subtract those also.</li>
	<li>Calculate enzyme activity in &mu;moles hr-1 g dry mass-1 from the equation:</li>
</ol>
Enzyme activity = Final absorbance / (C x incubation time x sample dry mass)
Fluorescent Analysis of Extracellular Enzyme Activity in Natural Waters
1. Preparation of Substrate, Standard, and Buffer Solutions for Fluorometric Analyses of Enzyme Activity
<ol>
	<li>Prepare 200 &mu;M solutions of MUB-linked substrates (e.g. 4-MUB-&beta;-glucopyranoside, 4-MUB-phosphate, 4-MUB-N-acetyl-&beta;-D-glucosaminide) by dissolving the appropriate substrate in sterile (autoclaved) distilled H2O in sterile 15 ml or 50 ml centrifuge tubes. Wrap tubes in aluminum foil to exclude light and store in refrigerator prior to use. Substrates should be stable for at least 1 week if stored in this way.</li>
	<li>Prepare a MUB standard by making a stock solution of 100 &mu;M 4-methylumbelliferone in sterile distilled H2O. Store refrigerated in amber or foil-wrapped bottle. Immediately prior to use, dilute the 100 &mu;M stock solution by 1/10 into sterile H2O to make a working solution of 10 &mu;M for enzyme assays.</li>
	<li>Prepare a stock solution of 100 mM bicarbonate buffer by dissolving 8.4 g of NaHCO3 into 1 L H2O and autoclaving. Dilute this stock solution 1/20 into sterile H2O as needed to make a working solution of 5 mM for enzyme assays.</li>
</ol>
2. Organizing Water Samples on a 96-Well Black Microplate
<ol>
	<li>Organize a microplate for each enzyme following the example shown in Figure 2. Note that allowing for adequate replication, standards, and controls, this procedure can assay the activity of a single enzyme for up to nine water samples on one 96-well black microplate.</li>
	<li>Pour approximately 5 ml of the first sample into a pipette reservoir and use an 8-channel pipettor to pipette200 &mu;l into all of the wells in column 1 of the microplate(s). Discard used pipette tips and repeat as needed for each water sample to fill columns 1-9.</li>
</ol>
3. Setting up Sample, Standard, Quench, and Substrate Controls
<ol>
	<li>Set up controls to account for samples, standards, substrate and quenching on the same black microplate as the samples (Figure 2).</li>
	<li>Sample controls contain sample water and bicarbonate buffer, and are not used in the activity calculations but will demonstrate reading consistency throughout the course of the experiment. The quench controls consist of sample water and a standard amount of the fluorescent tag and are used to measure the diffraction of fluorescence in sample water. Substrate and standard controls are made up of either substrate-linked or the standard fluorescent tag, respectively, and bicarbonate buffer.</li>
	<li>Pour approximately 5 ml of 5 mM bicarbonate buffer into a clean pipette reservoir. Pipette 50 &mu;l of buffer into microplate wells 1 through 9 in Rows D and E to form two replicate wells of sample controls per sample. Change pipette tips then transfer 200 &mu;l of bicarbonate buffer to wells 10 through 12 in Rows A and H.</li>
	<li>Reduce ambient lighting by dimming or turning off lights as the fluorescent standard is light sensitive.</li>
	<li>Pour approximately 5 ml of 10 &mu;M 4-methylumbelliferone into a clean pipette reservoir. Pipette 50 &mu;l into microplate wells 1 through 12 in Row H, and into wells 1 through 9 into Rows G and F to form three replicates of quench controls per sample and overall standard controls. Either place the microplate in the dark or cover with an opaque lid to reduce light degradation of MUB.</li>
	<li>Turn on the fluorometer and set-up any necessary software to be ready to read before adding the substrate. Note: some fluorometer bulbs may require a warm-up time of 3 min or more.</li>
	<li>Pour approximately 5 ml of the appropriate MUB-linked substrate (e.g. 4-MUB-phosphate) into a clean pipette reservoir. Use a 12-channel pipettor to pipette 50 &mu;l into microplate wells 1 through 12 in Row A, and into wells 1 through 9 in Rows B and C to form three replicate assays for each sample and three substrate controls. Immediately proceed to step 4.1, recording fluorescence.</li>
</ol>
4. Recording Fluorescence
<ol>
	<li>Read the initial fluorescence immediately after substrate addition to the microplate. After reading fluorescence, incubate the microplate at RT (22 &deg;C) either in the dark or covered with an opaque lid to reduce light degradation of MUB.</li>
	<li>The incubation time required to measure the maximum potential enzyme activity in a water sample will depend on the enzyme concentration within the sample. Since this is unknown before the assay is completed, the microplate will have to be read at multiple time steps. Typically, reading at intervals of 10-15 min over the course of 1 hr is acceptable for many enzymes, although samples with very high activity for certain enzymes may peak before 10 min.</li>
	<li>Continue reading fluorescence in the microplate at your designated intervals for at least 1 hr. Be sure to keep the microplate covered or in the dark between readings.</li>
</ol>
5. Calculation of Enzyme Activity per Volume of Water
<ol>
	<li>For each sample at each time interval calculate the: mean initial sample fluorescence (wells D and E), the mean final sample fluorescence (wells A-C), the mean standard fluorescence (wells H10-11), and the mean quench control fluorescence (wells F-G).</li>
	<li>For each time interval, calculate enzyme activity in nmoles hr-1 ml-1 from the equation:</li>
</ol>
Enzyme activity = (mean sample fluorescence - mean initial sample fluorescence) / ((mean standard fluorescence / 0.5 mol) x (mean quench control fluorescence / mean standard fluorescence) x (0.2 ml) x (time in hr))
<ol start="3">
	<li>Examine the activity values calculated for each time step. Determine final potential activity from the time step with the highest activity. If activity values continue to increase then later time steps may be required; if activity values fall throughout the course of the run, then run again with shorter time steps. Final activity is in nmoles of substrate consumed hr-1 ml-1 but can be scaled up to express as &mu;moles hr-1 L-1.</li>
</ol>

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K., Ochs, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+population+dynamics+of+freshwater+armored+dinoflagellates+in+a+small+lake+in+Mississippi.">The population dynamics of freshwater armored dinoflagellates in a small lake in Mississippi.</a> Journal of freshwater ecology. 20, 617-626 (2005).</li><li>Sinsabaugh, R. L., Lauber, C. L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stoichiometry+of+soil+enzyme+activity+at+global+scale.">Stoichiometry of soil enzyme activity at global scale.</a> Ecology letters. 11, 1252-1264 (2008).</li><li>Jackson, C. R., Foreman, C. M., Sinsabaugh, R. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbial+enzyme+activities+as+indicators+of+organic+matter+processing+rates+in+a+Lake+Erie+coastal+wetland.">Microbial enzyme activities as indicators of organic matter processing rates in a Lake Erie coastal wetland.</a> Freshwater biology. 34, 329-342 (1995).</li><li>Jackson, C. R., Vallaire, S. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbial+activity+and+decomposition+of+fine+particulate+organic+matter+in+a+Louisiana+cypress+swamp.">Microbial activity and decomposition of fine particulate organic matter in a Louisiana cypress swamp.</a> Journal of the north american benthological society. 26, 743-753 (2007).</li><li>Jackson, C. R., Liew, K. C., Yule, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Structural+and+functional+changes+with+depth+in+microbial+communities+in+a+tropical+Malaysian+peat+swamp+forest.">Structural and functional changes with depth in microbial communities in a tropical Malaysian peat swamp forest.</a> Microbial ecology. 57, 402-412 (2009).</li><li>Rietl, A. J., Jackson, C. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+the+ecological+restoration+practices+of+prescribed+burning+and+mechanical+thinning+on+soil+microbial+enzyme+activities+and+leaf+litter+decomposition.">Effects of the ecological restoration practices of prescribed burning and mechanical thinning on soil microbial enzyme activities and leaf litter decomposition.</a> Soil biology and biochemistry. 50, 47-57 (2012).</li><li>Smart, K. A., Jackson, C. 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The authors declare that they have no competing financial interests.

Determining the activity of a variety of microbial extracellular enzymes in soils and sediments can provide useful insights into rates of nutrient mineralization and organic matter processing17. However, soils can vary in their moisture levels, so it is important to standardize activity to soil dry weight. This requires an additional drying step (typically of two days) beyond simply measuring enzyme activity. Thus, in contrast to assays of enzyme activity in water samples that provide near instantaneous result...

Microorganisms such as bacteria and fungi obtain nutrients and carbon from complex organic compounds through the production of extracellular enzymes. These enzymes typically hydrolyze polymers into smaller subunits that can be taken into the cell. Therefore, at an ecological level, these microbial extracellular enzymes are responsible for much of the nutrient mineralization and organic matter decomposition that occurs in natural environments. Enzymes such as cellobiohydrolase (CBH) and &beta;-glucosidase are important for cellulose degradation and work in unison to catalyze the hydrolysis of cellulose to glucose1,2, which provides a utilizable carbon substrate for microbial uptake and assimilation. The enzyme phosphatase releases soluble inorganic phosphate groups from organophosphates, essentially mineralizing phosphate and making it available for use by most organisms3. Other enzymes, such as N-acetylglucosaminidase (NAGase), are important in chitin degradation and can make both carbon and nitrogen available for microbial acquisition4.
One of the procedures for the assay of microbial extracellular enzyme activity in natural environments is the use of artificial p-nitrophenyl (pNP) linked substrates, an approach that was originally developed to detect soil phosphatase activity5. This approach relies on the detection of a colored end product, p-nitrophenol, which is released when the artificial substrate is hydrolyzed by the appropriate enzyme. The p-nitrophenol can be subsequently quantified colorimetrically by measuring its absorbance at around 400-410 nm. This method has since been applied to detect other enzymes such as NAGase6, and has been used in various studies looking at microbial extracellular enzyme activity in soils and sediments7-9.
An alternative approach that was originally developed to assess extracellular glucosidase activity in aquatic environments10,11 makes use of 4-methylumbelliferone (MUB) linked substrates. The end product released (4-methylumbelliferone) is highly fluorescent and can be detected using a fluorometer with an excitation/emission setting around 360/460 nm. A variety of MUB-linked artificial substrates are available, permitting the fluorometric measurement of the activity of at least as many enzymes (e.g. &beta;-glucosidase, cellobiohydrolase, NAGase, phosphatase) as can be assayed using the pNP-substrate colorimetric procedure. Other microbial extracellular enzymes, such as the protein-degrading leucine aminopeptidase, can be assayed fluorometrically using 7-amino-4-methylcoumarin (COU) linked substrates. Both MUB- and COU-linked substrates have been used to determine enzyme activity in various terrestrial and aquatic samples12,13.
While previous studies have described fluorometric or colorimetric microplate approaches to determine extracellular enzyme activity14; there is a need for a clear presentation of how to conduct such assays. Here we demonstrate procedures for conducting high throughput microplate techniques for the analysis of extracellular enzyme activity in soils and sediments using the colorimetric pNP-linked substrates approach and in natural waters using the fluorescent MUB-linked substrates technique. We focus on the measurement of the activities of &beta;-glucosidase, NAGase, and phosphatase as these enzymes can be tied to carbon, nitrogen, and phosphorus cycling, respectively. However, the procedures described here can be applied to the measurement of other extracellular enzymes using different artificial substrates.

<table border="1">
		<tbody>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>REAGENTS AND MATERIALS</td>
		 </tr>
		 <tr>
			<td>Glacial acetic acid</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>Sodium acetate</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>Sodium hydroxide</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>p-Nitrophenol</td>
			<td>Fisher</td>
			<td>BP612-1</td>
			<td>Alternates available</td>
		 </tr>
		 <tr>
			<td>p-Nitrophenyl (pNP)-phosphate</td>
			<td>Sigma</td>
			<td>N3234</td>
			<td>pNP-substrate</td>
		 </tr>
		 <tr>
			<td>pNP-&beta;-glucopyranoside</td>
			<td>Sigma</td>
			<td>N7006</td>
			<td>pNP-substrate</td>
		 </tr>
		 <tr>
			<td>pNP-&beta;-N-acetylglucosaminide</td>
			<td>Sigma</td>
			<td>N9376</td>
			<td>pNP-substrate</td>
		 </tr>
		 <tr>
			<td>Clear 96-well microplates</td>
			<td>Fisher</td>
			<td>12-563-301</td>
			<td>Alternates available</td>
		 </tr>
		 <tr>
			<td>96-well deep well blocks</td>
			<td>Costar</td>
			<td>3958</td>
			<td>Alternates available</td>
		 </tr>
		 <tr>
			<td>Aluminum weigh pans</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>Sterile 15 ml centrifuge tubes</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>Sterile 50 ml centrifuge tubes</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>4-Methylumbelliferone</td>
			<td>Sigma</td>
			<td>M1381</td>
			<td></td>
		 </tr>
		 <tr>
			<td>4-Methylumbelliferyl (MUB)-phosphate</td>
			<td>Sigma</td>
			<td>M8883</td>
			<td>MUB-substrate</td>
		 </tr>
		 <tr>
			<td>4-MUB-glucopyranoside</td>
			<td>Sigma</td>
			<td>M3633</td>
			<td>MUB-substrate</td>
		 </tr>
		 <tr>
			<td>4-MUB-N-acetylglucosaminide</td>
			<td>Sigma</td>
			<td>M2133</td>
			<td>MUB-substrate</td>
		 </tr>
		 <tr>
			<td>Sodium bicarbonate</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td>Black 96-well microplate</td>
			<td>Costar</td>
			<td>3792</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Pipette reservoir</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		 <tr>
			<td></td>
			<td></td>
			<td></td>
			<td>EQUIPMENT</td>
		 </tr>
		 <tr>
			<td>Centrifuge</td>
			<td>Eppendorf</td>
			<td>5810R</td>
			<td></td>
		 </tr>
		 <tr>
			<td>Centrifuge rotor</td>
			<td>Eppendorf</td>
			<td>A-4-81</td>
			<td>For microplates/deep-well blocks</td>
		 </tr>
		 <tr>
			<td>Microplate reader</td>
			<td>BioTek</td>
			<td>Synergy HT</td>
			<td>Alternates available</td>
		 </tr>
		 <tr>
			<td>Microplate fluorometer</td>
			<td>BioTek</td>
			<td>FLx 800</td>
			<td>Alternates available</td>
		 </tr>
		 <tr>
			<td>8-channel pipettor</td>
			<td></td>
			<td></td>
			<td>Various suppliers</td>
		 </tr>
		</tbody>
 </table>

determination of microbial extracellular enzyme activity in waters, soils, and sediments using high throughput microplate assays

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

Soils and aquatic sediments typically have appreciable levels of extracellular enzyme activity as a result of attached microbial communities (biofilms) growing on the surface of particles. Figure 3 shows how this activity changes depending on the size of particles obtained from the surface sediment of a third order stream in northern Mississippi, USA. A previous study has shown that the bacterial communities on sediment particles from this stream can be separated into three distinct groups based on molec...

Watch this Scientific Journal Video about Determination of Microbial Extracellular Enzyme Activity in Waters, Soils, and Sediments using High Throughput Microplate Assays at JoVE.com

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

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

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

determination-microbial-extracellular-enzyme-activity-waters-soils

Research

JoVE Journal

Environment

38.7K Views.  The University of Mississippi. Microplate based procedures are described for the colorimetric or fluorometric analysis of extracellular enzyme activity. These procedures allow for the rapid assay of such activity in large numbers of environmental samples within a manageable time frame.

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

High-throughput Fluorometric Measurement of Potential Soil Extracellular Enzyme Activities

Microbes in soils and other environments produce extracellular enzymes to depolymerize and hydrolyze organic macromolecules so that they can be assimilated for energy and nutrients. Measuring soil microbial enzyme activity is crucial in understanding soil ecosystem functional dynamics. The general concept of the fluorescence enzyme assay is that synthetic C-, N-, or P-rich substrates bound with a fluorescent dye are added to soil samples. When intact, the labeled substrates do not fluoresce. Enzyme activity is measured as the increase in fluorescence as the fluorescent dyes are cleaved from their substrates, which allows them to fluoresce. Enzyme measurements can be expressed in units of molarity or activity. To perform this assay, soil slurries are prepared by combining soil with a pH buffer. The pH buffer (typically a 50 mM sodium acetate or 50 mM Tris buffer), is chosen for the buffer&#39;s particular acid dissociation constant (pKa) to best match the soil sample pH. The soil slurries are inoculated with a nonlimiting amount of fluorescently labeled (i.e. C-, N-, or P-rich) substrate. Using soil slurries in the assay serves to minimize limitations on enzyme and substrate diffusion. Therefore, this assay controls for differences in substrate limitation, diffusion rates, and soil pH conditions; thus detecting potential enzyme activity rates as a function of the difference in enzyme concentrations (per sample).
Fluorescence enzyme assays are typically more sensitive than spectrophotometric (i.e. colorimetric) assays,&#160;but can suffer from interference caused by impurities and the instability of many fluorescent compounds when exposed to light; so caution is required when handling fluorescent substrates. Likewise, this method only assesses potential enzyme activities under laboratory conditions when substrates are not limiting. Caution should be used when interpreting the data representing cross-site comparisons with differing temperatures or soil types, as in situ soil type and temperature can influence enzyme kinetics.

To measure potential rates of soil extracellular enzyme activities, synthetic substrates that are bound to a fluorescent dye are added to soil samples. Enzyme activity is measured as the fluorescent dye is released from the substrate by an enzyme-catalyzed reaction, where higher fluorescence indicates more substrate degradation.

Microbes in soils and other environments produce extracellular enzymes to depolymerize and hydrolyze organic macromolecules so that they can be ...

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

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

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

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

Measurement of Greenhouse Gas Flux from Agricultural Soils Using Static Chambers

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

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

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

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 ...

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is built and can be transported to a temporary work space close to field sampling site(s) to facilitate rapid analysis. Cores are extruded into a portable glove bag, where they are sectioned and each 1-3 cm thick section (depending on core diameter) is sealed into 50 ml centrifuge tubes. Pore waters are separated with centrifugation outside of the glove bag and then returned to the glove bag for separation from the sediment. These extracted pore water samples can be analyzed immediately. Immediate analyses of redox sensitive species, such as sulfide, iron speciation, and arsenic speciation indicate that oxidation of pore waters is minimal; some samples show approximately 100% of the reduced species, e.g. 100% Fe(II) with no detectable Fe(III). Both sediment and pore water samples can be preserved to maintain chemical species for further analysis upon return to the laboratory.

A protocol for sectioning sediment cores and extracting pore waters under anoxic conditions in order to permit analysis of redox sensitive species in both solids and fluids is presented.

We demonstrate a method for sectioning sediment cores and extracting pore waters while maintaining oxygen-free conditions. A simple, inexpensive system is ...

Clean Sampling and Analysis of River and Estuarine Waters for Trace Metal Studies

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination control. Developments of some techniques aiming to reduce trace metal contamination in the last couple of decades have resulted in concentrations reported now being orders of magnitude lower than those in the past. These low concentrations often necessitate preconcentration of water samples prior to instrumental analysis of samples. Since contamination can appear in all phases of trace metal analyses, including sample collection (and during preparation of sampling containers), storage and handling, pretreatments, and instrumental analysis, specific care needs to be taken in order to reduce contamination levels at all steps. The effort to develop and utilize "clean techniques" in trace metal studies allows scientists to investigate trace metal distributions and chemical and biological behavior in greater details. This advancement also provides the required accuracy and precision of trace metal data allowing for environmental conditions to be related to trace metal concentrations in aquatic environments.
This protocol that is presented here details needed materials for sample preparation, sample collection, sample pretreatment including preconcentration, and instrumental analysis. By reducing contamination throughout all phases mentioned above for trace metal analysis, much lower detection limits and thus accuracy can be achieved. The effectiveness of "clean techniques" is further demonstrated using low field blanks and good recoveries for standard reference material. The data quality that can be obtained thus enables the assessment of trace metal distributions and their relationships to environmental parameters.

Special care using "clean techniques" is required to properly collect and process water samples for trace metal studies in aquatic environments. A protocol for sampling, processing, and analytical procedures with the aim of obtaining reliable environmental monitoring data and results with high sensitivity for detailed trace metal studies is presented.

Most of the trace metal concentrations in ambient waters obtained a few decades ago have been considered unreliable owing to the lack of contamination ...

A Bending Test for Determining the Atterberg Plastic Limit in Soils

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

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

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

Extraction and Analysis of Microbial Phospholipid Fatty Acids in Soils

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

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

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

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

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

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

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

Preparation of Authigenic Pyrite from Methane-bearing Sediments for In Situ Sulfur Isotope Analysis Using SIMS

Different sulfur isotope compositions of authigenic pyrite typically result from the sulfate-driven anaerobic oxidation of methane (SO4-AOM) and organiclastic sulfate reduction (OSR) in marine sediments. However, unravelling the complex pyritization sequence is a challenge because of the coexistence of different sequentially formed pyrite phases. This manuscript describes a sample preparation procedure that enables the use of secondary ion mass spectroscopy (SIMS) to obtain in situ &#948;34S values of various pyrite generations. This allows researchers to constrain how SO4-AOM affects pyritization in methane-bearing sediments. SIMS analysis revealed an extreme range in &#948;34S values, spanning from -41.6 to +114.8&#8240;, which is much wider than the range of &#948;34S values obtained by the traditional bulk sulfur isotope analysis of the same samples. Pyrite in the shallow sediment mainly consists of 34S-depleted framboids, suggesting early diagenetic formation by OSR. Deeper in the sediment, more pyrite occurs as overgrowths and euhedral crystals, which display much higher SIMS &#948;34S values than the framboids. Such 34S-enriched pyrite is related to enhanced SO4-AOM at the sulfate-methane transition zone, postdating OSR. High-resolution in situ SIMS sulfur isotope analyses allow for the reconstruction of the pyritization processes, which cannot be resolved by bulk sulfur isotope analysis.

Preparation of Authigenic Pyrite from Methane-bearing Sediments for In Situ Sulfur Isotope Analysis Using SIMS

Analyses of the sulfur isotopic composition (&#948;34S) of pyrite from methane-bearing sediments have typically focused on bulk samples. Here, we applied secondary ion mass spectroscopy to analyze the &#948;34S values of various pyrite generations to understand the diagenetic history of pyritization.