This research was partly supported by the National Science Foundation&#39;s RAPID program (NSF-1048925, 1048919, and 1048914) to Alison Keimowitz, Ming-Kuo Lee, Benedict Okeke, and James Saunders.

1. Preparation of Equipment
<ol>
	<li>Preparation of Core Liners
	<ol>
		<li>Calculate thickness of core slice that will be obtained using Volume = &#960;r2 x thickness; the final volume must be &#60;50 cm3. With a 10 cm core diameter, 2 cm thick slices may be obtained. 
		NOTE: It is not necessary to have the volume be a full 50 ml, but porewater volumes obtained will be proportionately smaller.</li>
		<li>Using a jigsaw (or similar) slice one core liner, or a plastic tube of identical diameter, into 2 cm rings (or other thickness if using a different diameter core). Obtain 3-5 rings.</li>
		<li>Clean all plastic material that will come in contact with sediment, including core liners, core caps, rings, core slicers, centrifuge tubes, syringes, and disposable spoons. (Soak plastic materials in 10% HCl for 24 hr, rinse plastic materials 3x in nanopure (22 M&#937;) water, and allow materials to air dry, preferably in a laminar flow hood, before packing.)</li>
	</ol>
	</li>
	<li>Preparation of Laboratory Jack
	<ol>
		<li>Cut a piece of plywood, using a jigsaw, to cover the 6&#34; x 6&#34; size of the top plate of the laboratory jack and drill a hole in the center of this plywood using a hole saw attachment to a drill. 
		NOTE: This hole should be just slightly larger than the diameter of PVC extender pieces; for the sizes used here, the hole should be 2 &#190;&#34;.</li>
		<li>Drill four small holes in the edges of this plywood to match the holes on the top plate of the laboratory jack using a regular drill bit on a hand drill or drill press. Fasten the plywood to the laboratory jack with zip ties. 
		NOTE: There should be no &#34;wiggle&#34; in this plywood.</li>
	</ol>
	</li>
	<li>Drill a hole in the center of an approximately 2&#39; x 1.5&#39; plywood slightly larger than the outer diameter of the core liners using a hole saw. For 10 cm diameter core liners, a 10.5-11.5 cm hole will be appropriate. This wood is the core guidance plate.</li>
	<li>Preparation of Core Extruder (Plunger)
	<ol>
		<li>Obtain a rubber laboratory stopper that fits snugly inside the core liners. If none is available, shave one to size using a razor. Do not use a too-small stopper.</li>
		<li>Screw the laboratory stopper to an approximately 1&#39;-1.5&#39; long, 1&#34; diameter dowel or broom handle. Cover the screw head on the face of the stopper with waterproof electrical tape. 
		NOTE: The larger side of the stopper should face away from the dowel.</li>
	</ol>
	</li>
	<li>Cut an approximately 1.5&#39; length of PVC pipe into ~6&#34; long sections. 
	NOTE: The internal diameter should be larger than the dowel in step 1.4.2, but smaller than the stopper in step 1.4.1. This protocol assumes using standard 2&#34; PVC with a true outer diameter of 2.375&#34;. These PVC pieces are the core extenders.</li>
	<li>Assemble all other needed equipment listed in the Materials List. 
	NOTE: This should be gathered in the home laboratory and brought to the field laboratory.</li>
</ol>
2. Setting up the Field Laboratory Station
<ol>
	<li>Setting up the Glove Bag
	<ol>
		<li>Clamp the core guidance plate to the work surface (countertop, lab surface, etc.). Ensure that the hole for the core liner is over the work surface, but is open.</li>
		<li>Place the disposable glove bag over the core guidance plate, and run the tubing from the tubing entry of the glove bag to the regulator of the N2 tank. Make sure the tank is safely secured using the cylinder bench clamp.</li>
		<li>Tape the tubing to the glove bag to seal up this entry point by encircling the bag around the exterior of the tubing with electrical tape. Ensure that about 8&#34; of tubing extends into the interior of the glove bag. Slide the tubing clamp onto the tubing inside the glove bag; leave this clamp open.</li>
		<li>Cut an &#34;X&#34; in the bottom of the glove bag over the hole in the core guidance plate using a box cutter or straight razor. This x should be smaller than the core liner diameter.</li>
		<li>Load the glove bag with the items found in Table 1.</li>
	</ol>
	</li>
	<li>Setting up the Core
	<ol>
		<li>Place the laboratory jack on the floor underneath the work area where the glove bag is affixed. Place the core through the &#34;X&#34; cut in the bottom of the glove bag in step 2.1.4, keeping it in an upright position. 
		NOTE: About 4-6&#34; of the core should extend above the core stabilization plate.</li>
		<li>Hold the core steady to allow researcher 2 to perform steps 2.2.3 through 2.2.6.</li>
		<li>Tape the core to the plastic glove bag around the &#34;X&#34;, using plenty of good sealing electrical tape.</li>
		<li>Insert the handle of the core extruder into a 2&#34; diameter, ~6&#34; long PVC spacer, followed by a ~3&#34; long coupling, followed by another spacer. Continue this pattern until the PVC entirely covers the handle; it may extend a short distance past the end of the handle. Place the core extruder (with PVC spacers and couplings) underneath the bottom of the core.</li>
		<li>Support the core extruder with the laboratory jack so that the extruder can support the core. At this time, keep the jack as low as possible and use core spacers wherever possible.</li>
		<li>Use a box cutter to cut carefully around the bottom core cap. This cut should result in a ring of core cap being left around the outside of the core liner and a flat circular portion of the core cap in place against the core materials.</li>
		<li>Insert the hands into the glove bag (researcher 1). Hold the core steady so that it does not move further above the core stabilization plate while researcher 2 begins to turn the laboratory jack slowly to raise the core. The core extruder should enter the core liner and begin to push the sediment upwards like a push pop.</li>
		<li>Wiggle the core extruder side to side slightly if needed to insert it into the core liner (researcher 2). Be prepared for a slight pop when it enters.</li>
		<li>Continue to raise the core until the top of the cored material is at or near the top of the core liner (researchers 1 and 2). Note that the core cap should remain atop the core during this procedure.</li>
	</ol>
	</li>
	<li>Sealing the Glove Bag
	<ol>
		<li>Double check that all necessary supplies are in the glove bag; turn on the portable oxygen meter.</li>
		<li>Open all centrifuge tubes, water bottles, and other items that contain trapped air. If the core has headspace above the standing water, open the top cap to purge this headspace.</li>
		<li>Turn on the regulator so that the flow of nitrogen through the tube into the glove bag is at a moderate rate. Generally a pressure of ~15 psi at the last regulator stage with all valves open is appropriate. 
		NOTE: This should be fast enough to feel like a strong breeze on skin, but not so strong that it scatters glove bag supplies.</li>
		<li>Point the nitrogen into all areas of the glove bag, and push the nitrogen out the main opening to the best of one&#39;s ability. Turn off the nitrogen.</li>
		<li>Purge the glove bag three times by repeating steps 2.3.6 through 2.3.9 three times. 
		NOTE: One person can accomplish this, but it may be easier for two people so that one can keep his or her hands in the gloves that are a part of the glove bag.</li>
		<li>Seal the main opening of the glove bag by wrapping 1-2 bungee cords around it several times. Turn on the nitrogen to a similar flow as in 2.3.3. Place researcher 1&#39;s hands in the built-in gloves of the glove bag.</li>
		<li>Move the nitrogen tube from one area of the bag to another to fill the glove bag with nitrogen. Point the tube into any crevices such as opened centrifuge tube, above the water in a squirt bottle, etc.</li>
		<li>Turn off the nitrogen by closing the tubing clamp. Note that this can be accomplished without removing researcher 1&#39;s hands from the glove bag.</li>
		<li>Open the front of the glove bag and using researcher 1&#39;s body, push as much gas out of the bag as possible. 
		NOTE: The bag should be flat around the supplies inside it at this point.</li>
		<li>Fill the glove bag to a comfortable pressure and turn off the nitrogen using the tubing clamp. Some trial and error may be required to find a comfortable pressure, as too full and it is difficult to move one&#39;s arms, while too empty and it is difficult to see and manipulate objects. Open the nitrogen tubing clamp for short periods to refill the bag if it seems to have a slow leak or is getting emptier for any reason. Open the front of the bag to let out small amounts of nitrogen if it becomes uncomfortably full.</li>
		<li>Check the level of dissolved oxygen on the portable oxygen meter. It should be below 1%.</li>
		<li>Insert the glove bag gloves into disposable gloves. These will improve dexterity. Change them whenever they get dirty or torn; discard in the in-bag waste container.</li>
	</ol>
	</li>
</ol>
3. Sectioning the Core
NOTE: this portion of the procedure is much more easily accomplished with two researchers.
<ol>
	<li>Remove standing water using a syringe. Syringe filter this water and place into 50 ml centrifuge tubes.</li>
	<li>Remove a Core Section
	<ol>
		<li>Place a core sectioning ring above the top of the core liner. Raise the core up (section 3.3) until the top of the sediment is at the top of the ring.</li>
		<li>Insert the core slicer between the top of the core liner and the ring. The sediment section is now sitting atop the core slicer.</li>
		<li>Move the sediment to a 50 ml tube using the disposable spoons. Cap the tube tightly once it is full.</li>
		<li>Rinse the core slicer and core sectioning ring with the nanopure water; squirt the dirty waste into the waste container inside the glove bag. Disposable spoons can also be rinsed, or discarded, depending on the number needed. Dry the core slicer and core sectioning ring with paper towels.</li>
		<li>Repeat step 3.2 until no further core material remains.</li>
	</ol>
	</li>
	<li>Raising the Core
	<ol>
		<li>Throughout 3.2 it is necessary to raise the core. Do this in small increments using the laboratory jack. Perform the following three steps once the laboratory jack is fully extended.</li>
		<li>Hold the core in place in the glove bag (researcher 1).</li>
		<li>Lower the laboratory jack to its lowest setting, when it is fully compressed (researcher 2). Ensure that the core is held in place with researcher 1&#39;s hands within the glove bag and researcher 2&#39;s hands supporting from beneath.</li>
		<li>Fill the space between the bottom end of the core extruder and the jack with PVC spacers and fittings. Ensure that the core is securely supported by the core extender before letting go of the top of the core.</li>
	</ol>
	</li>
</ol>
4. Extracting Porewaters
<ol>
	<li>Open the glove bag and remove the rack of centrifuge tubes.</li>
	<li>Clean up the glove bag as needed by removing any soil, liquid, or condensation by wiping out the inside with paper towels, water, etc. Empty waste container.</li>
	<li>Reseal the glove bag loosely with bungee cords.</li>
	<li>Weigh centrifuge tubes which now contain sectioned sediment (if desired).</li>
	<li>Wrap the tops of the centrifuge tubes with electrical tape to seal the cap/tube junction.</li>
	<li>Balance the tubes for centrifugation by weighing them after taping. Add or remove electrical tape to get weights within 0.5 g. 
	NOTE: Generally, tubes from the top of the core will be lighter than those from the bottom.</li>
	<li>Place tubes in centrifuge and centrifuge at maximum acceleration (1,100 x g is recommended) for 20 min. 
	NOTE: Higher centrifugation rates will permit greater separation of porewaters.</li>
	<li>Remove electrical tape from the centrifuge tubes either by unpeeling or by slicing with a razor prior to returning the tubes to the glove bag. Return centrifuge tubes to rack after centrifugation and return to the glove bag.</li>
	<li>Add another set of centrifuge tubes, syringes, and syringe filters to the glove bag. These new tubes will hold porewaters and can be pre-labeled.</li>
	<li>Purge the glove bag again as in step 2.3.5. If the glove bag was only opened briefly to remove the tubes, and was not cleaned, it will be acceptable to purge it only two times. If an oxygen monitor is being used, ensure that the atmosphere in the glove bag is &#60;1% O2. Do not forget to open the newly added centrifuge tubes and purge them.</li>
	<li>Insert hands to the gloves in the glove bag; as before, cover with disposable gloves.</li>
	<li>Open one tube. Remove the porewaters from above the sediment using a syringe, then attach a syringe filter to the tip of the syringe. If porewaters and sediments are well separated, porewaters may be poured directly into a syringe with the barrel removed and a syringe filter on the tip.</li>
	<li>Push the water through the syringe filter into the appropriate centrifuge tube. 
	NOTE: Some force will be needed; more than one syringe filter per sample may be required.</li>
	<li>Replace the caps on the centrifuge tube containing sediment and the tube containing porewaters.</li>
	<li>Repeat steps 4.13 through 4.15 for each sample.</li>
	<li>Open glove bag and remove samples. Both sediment and porewater tubes can be re-weighed.</li>
	<li>Analyze porewaters immediately if desired. Analyses can include (but are not limited to) ion chromatography for major ions11, ferrozine for iron speciation12, arsenic speciation by voltammetry13, and sulfide speciation14.</li>
	<li>Freeze porewaters and sediments in the dry shipper (approximately -80 &#176;C) to preserve speciation for later analyses if appropriate for the planned analyses. It may also be appropriate to keep samples cool in airtight containers or in nitrogen purged aluminum bags.</li>
	<li>Re-use the glove bag for a second core (starting at 2.2.2) if desired. After two cores, the glove bag usually needs to be changed.</li>
</ol>

<ol>
	<li>Chapman, P. M., Wang, F., Germano, J. D., Batley, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pore+water+testing+and+analysis:+the+good,+the+bad,+and+the+ugly.">Pore water testing and analysis: the good, the bad, and the ugly.</a> Mar Poll Bull. 44, 359-366 (2002).</li><li>Teasdale, P. R., Batley, G. E., Apte, S. C., Webster, I. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pore+water+sampling+with+sediment+peepers.">Pore water sampling with sediment peepers.</a> TrAC. 14, 250-256 (1995).</li><li>Steinmann, P., Shotyk, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chemical+composition,+pH,+and+redox+state+of+sulfur+and+iron+in+complete+vertical+porewater+profiles+from+two+Sphagnum+peat+bogs,+Jura+Mountains,+Switzerland.">Chemical composition, pH, and redox state of sulfur and iron in complete vertical porewater profiles from two Sphagnum peat bogs, Jura Mountains, Switzerland.</a> Geochim Cosmochim Acta. 61, 1143-1163 (1997).</li><li>Bufflap, S. E., Allen, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Sediment+pore+water+collection+methods+for+trace+metal+analysis:+A+review.">Sediment pore water collection methods for trace metal analysis: A review.</a> Wat Res. 29, 165-177 (1995).</li><li>Glew, J., Smol, J., Last, W., Last, W., Smol, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Chapter+5,+Sediment+Core+Collection+and+Extrusion.">Chapter 5, Sediment Core Collection and Extrusion.</a> Developments in Paleoenvironmental Research. , 73-105 (2001).</li><li>Jahnke, R. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simple,+reliable,+and+inexpensive+pore-water+sampler.">A simple, reliable, and inexpensive pore-water sampler.</a> L&amp;O. 33, 483-487 (1988).</li><li>Bufflap, S. E., Allen, H. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparison+of+pore+water+sampling+techniques+for+trace+metals.">Comparison of pore water sampling techniques for trace metals.</a> Wat Res. 29, 2051-2054 (1995).</li><li>Zheng, Y., Anderson, R. F., van Geen, A., Kuwabara, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Authigenic+molybdenum+formation+in+marine+sediments:+a+link+to+pore+water+sulfide+in+the+Santa+Barbara+Basin.">Authigenic molybdenum formation in marine sediments: a link to pore water sulfide in the Santa Barbara Basin.</a> Geochim Cosmochim Acta. 64, 4165-4178 (2000).</li><li>Keimowitz, A. R., 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=Arsenic+redistribution+between+sediments+and+water+near+a+highly+contaminated+source.">Arsenic redistribution between sediments and water near a highly contaminated source.</a> Env Sci &amp; Tech. 39, 8606-8613 (2005).</li><li>Natter, M., 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=Level+and+Degradation+of+Deepwater+Horizon+Spilled+Oil+in+Coastal+Marsh+Sediments+and+Pore-Water.">Level and Degradation of Deepwater Horizon Spilled Oil in Coastal Marsh Sediments and Pore-Water.</a> Env Sci &amp; Tech. 46, 5744-5755 (2012).</li><li>Jackson, P. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Ion chromatography. , (1990).</li><li>Stookey, L. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ferrozine+-+A+New+Spectrophotometric+Reagent+For+Iron.">Ferrozine - A New Spectrophotometric Reagent For Iron.</a> Anal. Chem. 42, 779-781 (1970).</li><li>He, Y., Zheng, Y., Ramnaraine, M., Locke, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Differential+pulse+cathodic+stripping+voltammetric+speciation+of+trace+level+inorganic+arsenic+compounds+in+natural+water+samples.">Differential pulse cathodic stripping voltammetric speciation of trace level inorganic arsenic compounds in natural water samples.</a> Anal. Chim. Acta. 511, 55-61 (2004).</li><li>Cline, J. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spectrophotometric+Determination+of+Hydrogen+Sulfide+in+Natural+Waters.">Spectrophotometric Determination of Hydrogen Sulfide in Natural Waters.</a> L&amp;O. 14, 454-458 (1969).</li></ol>

The authors have nothing to disclose.

The technique described herein is a flexible one that can be adjusted for a wide range of locations, core sizes, core section thickness, etc. There are three essential components to this system.
First, prepare a core extrusion system of the right dimensions for the core to be analyzed. Instructions here are given assuming an approximately 30&#34; core; much longer cores may require more PVC extender pieces and PVC fittings to extrude fully. Plan the extrusion system and the packing carefully, as corrections in the field are much more difficult to manage.
Second, ensure that the glove bag is well purged and free of leaks or tears. The purpose of this protocol is to obtain porewaters in the same redox state in which they existed below ground. If oxidation occurs during pore sectioning or pore water extraction, the data obtained will not be usable.
Third, the centrifugation immediately allows separation of the porewaters from the sediments. If porewaters and sediments remain in contact after removal from the environment, reactions may continue and change. For example, if water flowing through the core was supplying nitrate, this would prevent the existing microbial community from using iron as a terminal electron acceptor; after removal of the core from the site, nitrate concentrations would begin to decrease and iron speciation could begin to change. Therefore rapid core sectioning and centrifugation allow the best &#34;snapshot&#34; to be taken.
Depending on the analyses desired, it may be desirable to weigh the centrifuge tubes before filling them with sediment and porewater. This will allow computation of the exact masses of porewater and sediment collected from each section. If this is not necessary, an average mass of the 50 ml centrifuge tubes may be assumed for each tube. This is usually sufficient. In general, a subsection of sediment may be removed, weighed, and dried again to obtain a percent dry mass value. When doing this, make sure to include the weight of porewaters removed as part of the calculation. Dried sediment may also be combusted to obtain a loss on ignition measurement.
It may be advisable to practice this procedure once or twice on a sample core before performing it on valuable field samples. Once it is mastered, however, this technique allows collection of pore waters and sediments from a wide range of environments in a straightforward, cost-effective way. The ability to maintain the in situ redox conditions allows a range of geochemical and geomicrobiological analyses on the collected samples.

Researchers often wish to study the redox state and geomicrobiology of a sediment-water system. This ideally utilizes data from both sediments and pore waters, as pore waters are often sensitive monitors of the system and are a common source, though not the only source, of ecological exposure to redox-sensitive heavy metals1 such as arsenic and uranium. Pore water data may be obtained in situ using diffusion equilibrium filters, also known as &#34;peepers,&#34; installed into the sediment2. Peepers are most commonly used in settings where the field site is known prior to beginning field work and where multiple visits over an extended period of time can be made to the field site, e.g. Shotyk3. Therefore many contexts do not permit the use of peepers, such as sites only accessible for a short time or where multiple exploratory samples are obtained to determine where further investigation should occur4. Additionally peepers do not sample sediment simultaneously to water sampling.
When it is desirable to sample sediment and water together, or in field sites where peeper installation is not feasible, the most common method to obtain sediment and water is sediment coring. Obtaining an unmixed core is a crucial precursor to the procedure described in this work5. Once a core is obtained pore waters may be obtained by squeezing6 or centrifugation; both have advantages and disadvantages. Centrifugation is generally considered the most reliable method for extracting porewaters from sediment cores,7 although care must be taken to prevent oxidation of sediments or pore waters.
In this method we describe core extrusion and centrifugation to extract pore waters with minimal oxidation. The authors have used the method described herein in a variety of contexts including marine8, contaminated lake9, and wetlands10. The representative data shown demonstrates that reducing conditions can be preserved. With the exception of the centrifuge, materials used are inexpensive, and this method can be applied to a wide variety of geochemical and geomicrobiological research questions.

<table border="0" cellpadding="0" cellspacing="0" width="811">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;Disposable glove bag(s).&#160;</td>
			<td>Sigma-Aldrich</td>
			<td>Z106089-1EA</td>
			<td>&#160;One per two cores to be processed is usually sufficient.</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">&#160;N2 tank</td>
			<td>Praxair</td>
			<td></td>
			<td>Often gas supply companies can deliver these directly to the field laboratory.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Nitrogen gas regulator</td>
			<td>VWR</td>
			<td>55850-478</td>
			<td>Or similar</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Several feet of tubing that fits the regulator</td>
			<td>VWR</td>
			<td>89403-862</td>
			<td>Or similar</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Safety equipment to secure the tank</td>
			<td>VWR</td>
			<td>60142-006</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Adjustable tubing clamp</td>
			<td>VWR</td>
			<td>62849-112</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Waterproof, good sealing electrical tape</td>
			<td>Scotch</td>
			<td>Super 33+</td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;2-4 short bungee cords</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Squirt bottles of nanopure water</td>
			<td>VWR</td>
			<td>16650-082</td>
			<td>Any similar bottle is fine; pack an additional supply of nanopure water to refill these.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Large supply of paper towels and kimwipes.</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">50 mL centrifuge tubes</td>
			<td>VWR</td>
			<td>21008-951</td>
			<td>Acid cleaned as described in protocol.&#160; At least 2/core section needed.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Several permanent in markers.</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;Several straight razor blades and box cutters.</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Centrifuge</td>
			<td>Beckman-Coulter</td>
			<td>Allegra X-22</td>
			<td>Faster rotor allows greater separation.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rotor to accommodate 50 mL tubes</td>
			<td>Beckman-Coulter</td>
			<td>SX-4250</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">]50 mL plastic syringes without black rubber tip on the barrel</td>
			<td>VWR</td>
			<td>66064-764&#160;</td>
			<td>Acid cleaned as described in protocol.&#160; At least 1/core section needed, plus 1 for overlying water.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Syringe filters compatible with aqueous solutions.</td>
			<td>VWR</td>
			<td>28143-310&#160;</td>
			<td>Either 0.45 &#956;m or 0.20 &#956;m poresizes may be used.&#160; Plan on five filters per core section processed.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plastic (disposable) spoons.</td>
			<td></td>
			<td></td>
			<td>Widely available; Acid cleaned as described in protocol.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Several boxes of disposable gloves.</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;">Large plastic beakers or other waste containers to place in the glove bag.</td>
			<td>VWR</td>
			<td>13890-148</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Laboratory balance</td>
			<td>VWR</td>
			<td>10205-008</td>
			<td>An available balance will be fine; high precision not required</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dry shipper, pre-charged with liquid nitrogen</td>
			<td>VWR</td>
			<td>82005-416</td>
			<td>Needed only if samples are being returned to the home laboratory for sensitive analyses.</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Laboratory notebooks</td>
			<td></td>
			<td></td>
			<td>Water repellent can be useful</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Core liners</td>
			<td>Watermark</td>
			<td>77280</td>
			<td>Available from Forrestry Suppliers</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Core caps</td>
			<td>Ben Meadows</td>
			<td>218105</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Core slicers</td>
			<td>McMaster Carr</td>
			<td>8707K111</td>
			<td>Cut this into 9 3x3 squares</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PVC spacers</td>
			<td>McMaster Carr</td>
			<td>48925K96</td>
			<td>Cut this into short lengths</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">PVC couplings</td>
			<td>McMaster Carr</td>
			<td>4880K76</td>
			<td>Approximately 12 needed</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dowel</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lab stopper</td>
			<td>VWR</td>
			<td>59580-400</td>
			<td>Check to ensure the correct size to fit snugly within the core liners</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Plywood for core guidance plate and top of lab jack</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lab jack</td>
			<td>VWR</td>
			<td>89260-826</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Clamps</td>
			<td></td>
			<td></td>
			<td>Widely available</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Portable oxygen monitor</td>
			<td>RKI instruments</td>
			<td>OX-07</td>
			<td></td>
		</tr>
	</tbody>
</table>

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.

The type of results obtained depends on analyses performed and on the geochemical setting from which the core was obtained. Dissolved oxygen may be measured in the extracted porewaters, but in many settings this will be zero below the first few cm of the core. Analyses that usually provide more meaningful information include iron speciation (Fe II/Fe III)12, arsenic speciation (As III/As V)13, and sulfide14. Presence of reduced species such as sulfide indicates both a reducing environment and that sufficient anoxia was maintained during core sectioning and pore water removal. Determination of other concentrations such as dissolved organic carbon, major ions, or trace metals is often performed on preserved samples upon return to the home laboratory. Geochemical gradients can generally be observed in pore waters, and maxima or minima of particular species may be seen at depth.
A core was taken in Bay Batiste, a wetland southeast of New Orleans, about nine months after the beginning of the Deepwater Horizon spill. This wetland was heavily oiled, and data obtained from the sediment core indicate high sulfide concentrations in the porewaters using a Hach Method (http://hach.com) based on14; see Figure 1. Maximum sulfide concentrations of 49.2 mg/L S2- are observed in the core section obtained between 24-27 cm depth. Total iron concentrations in these porewaters were consistently low (&#60;0.2 ppm) and no Fe(III) was detected.
<img alt="Figure 1" src="/files/ftp_upload/53393/53393fig1.jpg" /> 
Figure 1: Porewaters from Bay Batiste, Louisiana. Data shown is from porewaters extracted from a sediment core using the methods described herein; the core was obtained from Bayou Batiste, Louisiana, in the year following the Deepwater Horizon oil spill to the Gulf of Mexico. Dissolved sulfide concentrations in porewaters as a function of depth below the sediment-water interface. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/53393/53393fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Items to be loaded into glove bag at Step 2.1.7</td>
		</tr>
		<tr>
			<td>Waste containers</td>
		</tr>
		<tr>
			<td>Box of disposable gloves</td>
		</tr>
		<tr>
			<td>Kimwipes and paper towels</td>
		</tr>
		<tr>
			<td>Straight razors</td>
		</tr>
		<tr>
			<td>Squirt bottle(s) of dd H2O</td>
		</tr>
		<tr>
			<td>Permanent marker</td>
		</tr>
		<tr>
			<td>Disposable plastic spoons</td>
		</tr>
		<tr>
			<td>50 ml centrifuge tubesa in a rack; one per core section plus enough for overlying water.&#160;</td>
		</tr>
		<tr>
			<td>Syringe filters sufficient in number for filtering overlying water.&#160;</td>
		</tr>
		<tr>
			<td>Core liner rings</td>
		</tr>
		<tr>
			<td>Core slicers</td>
		</tr>
		<tr>
			<td>Portable Oxygen Meter</td>
		</tr>
		<tr>
			<td>a Plastic materials such as centrifuge tubes should be acid-cleaned according to instructions.</td>
		</tr>
	</tbody>
</table>
Table 1: Materials to seal into glove bag.

Watch this Scientific Journal Video about Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions at JoVE.com

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

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 ...

sediment-core-sectioning-extraction-pore-waters-under-anoxic

Nanyang Technological University

Research

JoVE Journal

Environment

8.9K Views.  Vassar College. 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.

Video: Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

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

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

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

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

A Protocol for Collecting and Constructing Soil Core Lysimeters

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

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

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

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 ...

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 ...

Reliable Method for Assessing Seed Germination, Dormancy, and Mortality under Field Conditions

We describe techniques for approximating seed bank dynamics over time using Helianthus annuus as an example study species. Strips of permeable polyester fabric and glue can be folded and glued to construct a strip of compartments that house seeds and identifying information, while allowing contact with soil leachate, water, microorganisms, and ambient temperature. Strips may be constructed with a wide range of compartment numbers and sizes and allow the researcher to house a variety of genotypes within a single species, different species, or seeds that have experienced different treatments. As opposed to individual seed packets, strips are more easily retrieved as a unit. While replicate packets can be included within a strip, different strips can act as blocks or can be retrieved at different times for observation of seed behavior over time. We used a high temperature glue gun to delineate compartments and sealed the strips once the seed and tags identifying block and removal times were inserted. The seed strips were then buried in the field at the desired depth, with the location marked for later removal. Burrowing animal predators were effectively excluded by use of a covering of metal mesh hardware cloth on the soil surface. After the selected time interval for burial, strips were dug up and seeds were assessed for germination, dormancy and mortality. While clearly dead seeds can often be distinguished from ungerminated living ones by eye, dormant seeds were conclusively identified using a standard Tetrazolium chloride colorimetric test for seed viability.

Here we present a protocol for assessing seed survivorship, germination and dormancy under field conditions using buried, labeled seed strips and tetrazolium chloride (TZ) viability testing.

We describe techniques for approximating seed bank dynamics over time using Helianthus annuus as an example study species. Strips of permeable polyester ...

Extraction of Organochlorine Pesticides from Plastic Pellets and Plastic Type Analysis

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment during manufacturing and transport. Because of their environmental persistence, they are widely distributed in the oceans and on beaches all over the world. They can act as a vector of potentially toxic organic compounds (e.g., polychlorinated biphenyls) and might consequently negatively affect marine organisms. Their possible impacts along the food chain are not yet well understood. In order to assess the hazards associated with the occurrence of plastic pellets in the marine environment, it is necessary to develop methodologies that allow for rapid determination of associated organic contaminant levels. The present protocol describes the different steps required for sampling resin pellets, analyzing adsorbed organochlorine pesticides (OCPs) and identifying the plastic type. The focus is on the extraction of OCPs from plastic pellets by means of a pressurized fluid extractor (PFE) and on the polymer chemical analysis applying Fourier Transform-InfraRed (FT-IR) spectroscopy. The developed methodology focuses on 11 OCPs and related compounds, including dichlorodiphenyltrichloroethane (DDT) and its two main metabolites, lindane and two production isomers, as well as the two biologically active isomers of technical endosulfan. This protocol constitutes a simple and rapid alternative to existing methodology for evaluating the concentration of organic contaminants adsorbed on plastic pieces.

Microplastics act as vector of potentially toxic organic contaminants with unpredictable effects. This protocol describes an alternative methodology for assessing the levels of organochlorine pesticides adsorbed on plastic pellets and identifying the polymer chemical structure. The focus is on pressurized fluid extraction and attenuated total reflectance Fourier transform infrared spectroscopy.

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment ...

Extraction and Characterization of Surfactants from Atmospheric Aerosols

Surface-active compounds, or surfactants, present in atmospheric aerosols are expected to play important roles in the formation of liquid water clouds in the Earth&#39;s atmosphere, a central process in meteorology, hydrology, and for the climate system. But because specific extraction and characterization of these compounds have been lacking for decades, very little is known on their identity, properties, mode of action and origins, thus preventing the full understanding of cloud formation and its potential links with the Earth&#39;s ecosystems.
In this paper we present recently developed methods for 1) the targeted extraction of all the surfactants from atmospheric aerosol samples and for the determination of 2) their absolute concentrations in the aerosol phase and 3) their static surface tension curves in water, including their Critical Micelle Concentration (CMC). These methods have been validated with 9 references surfactants, including anionic, cationic and non-ionic ones. Examples of results are presented for surfactants found in fine aerosol particles (diameter &#60;1 &#956;m) collected at a coastal site in Croatia and suggestions for future improvements and other characterizations than those presented are discussed.

Methods are presented for the targeted extraction of surfactants present in atmospheric aerosols and the determination of their absolute concentrations and surface tension curves in water, including their Critical Micelle Concentration (CMC).

Surface-active compounds, or surfactants, present in atmospheric aerosols are expected to play important roles in the formation of liquid water clouds in ...

Continuous Instream Monitoring of Nutrients and Sediment in Agricultural Watersheds

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in water resources is a prerequisite for understanding the drivers of the pollutant loads and for making informed water resource management decisions. The commonly used &#34;grab sampling&#34; method provides the concentrations of pollutants at the time of sampling (i.e., a snapshot concentration) and may under- or overpredict the pollutant concentrations and loads. Continuous monitoring of nutrients and sediment has recently received more attention due to advances in computing, sensing technology, and storage devices. This protocol demonstrates the use of sensors, sondes, and instrumentation to continuously monitor in situ nitrate, ammonium, turbidity, pH, conductivity, temperature, and dissolved oxygen (DO) and to calculate the loads from two streams (ditches) in two agricultural watersheds. With the proper calibration, maintenance, and operation of sensors and sondes, good water quality data can be obtained by overcoming challenging conditions such as fouling and debris buildup. The method can also be used in watersheds of various sizes and characterized by agricultural, forested, and/or urban land.

With the advancement of technology and the rise in end-user expectations, the need and use of higher temporal resolution data for pollutant load estimation has increased. This protocol describes a method for continuous in situ water quality monitoring to obtain higher temporal resolution data for informed water resource management decisions.

Pollutant concentrations and loads in watersheds vary considerably with time and space. Accurate and timely information on the magnitude of pollutants in ...

Microfluidic Devices for Characterizing Pore-scale Event Processes in Porous Media for Oil Recovery Applications

Microfluidic devices are versatile tools for studying transport processes at a microscopic scale. A demand exists for microfluidic devices that are resistant to low molecular-weight oil components, unlike traditional polydimethylsiloxane (PDMS) devices. Here, we demonstrate a facile method for making a device with this property, and we use the product of this protocol for examining the pore-scale mechanisms by which foam recovers crude oil. A pattern is first designed using computer-aided design (CAD) software and printed on a transparency with a high-resolution printer. This pattern is then transferred to a photoresist via a lithography procedure. PDMS is cast on the pattern, cured in an oven, and removed to obtain a mold. A thiol-ene crosslinking polymer, commonly used as an optical adhesive (OA), is then poured onto the mold and cured under UV light. The PDMS mold is peeled away from the optical adhesive cast. A glass substrate is then prepared, and the two halves of the device are bonded together. Optical adhesive-based devices are more robust than traditional PDMS microfluidic devices. The epoxy structure is resistant to swelling by many organic solvents, which opens new possibilities for experiments involving light organic liquids. Additionally, the surface wettability behavior of these devices is more stable than that of PDMS. The construction of optical adhesive microfluidic devices is simple, yet requires incrementally more effort than the making of PDMS-based devices. Also, though optical adhesive devices are stable in organic liquids, they may exhibit reduced bond-strength after a long time. Optical adhesive microfluidic devices can be made in geometries that act as 2-D micromodels for porous media. These devices are applied in the study of oil displacement to improve our understanding of the pore-scale mechanisms involved in enhanced oil recovery and aquifer remediation.

The goal of this procedure is to easily and rapidly produce a microfluidic device with customizable geometry and resistance to swelling by organic fluids for oil recovery studies. A polydimethylsiloxane mold is first generated, and then used to cast the epoxy-based device. A representative displacement study is reported.

Microfluidic devices are versatile tools for studying transport processes at a microscopic scale. A demand exists for microfluidic devices that are ...

Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only recently have microplastics in freshwater systems, such as rivers and lakes, been quantified. Microplastic sampling at the surface usually consists of deploying drift nets behind either a stationary or moving boat, which limits the sampling to environments with low levels of suspended sediments and floating or submerged debris. Previous studies that employed drift nets to collect microplastic debris typically used nets with &#8805;300 &#181;m mesh size, allowing plastic debris (particles and fibers) below this size to pass through the net and elude quantification. The protocol detailed here enables: 1) sample collection in environments with high suspended loads and floating or submerged debris and 2) the capture and quantification of microplastic particles and fibers &lt;300 &#181;m. Water samples were collected using a peristaltic pump in low-density polyethylene (PE) containers to be stored before filtering and analysis in the lab. Filtration was done with a custom-made microplastic filtration device containing detachable union joints that housed nylon mesh sieves and mixed cellulose ester membrane filters. Mesh sieves and membrane filters were examined with a stereomicroscope to quantify and separate microplastic particulates and fibers. These materials were then examined using a micro-attenuated total reflectance Fourier transform infrared spectrometer (micro ATR-FTIR) to determine microplastic polymer type. Recovery was measured by spiking samples using blue PE particulates and green nylon fibers; percent recovery was determined to be 100% for particulates and 92% for fibers. This protocol will guide similar studies on microplastics in high velocity rivers with high concentrations of sediment. With simple modifications to the peristaltic pump and filtration device, users can collect and analyze various sample volumes and particulate sizes.

Most microplastic research to date has occurred in marine systems where suspended solid levels are relatively low. Focus is now shifting to freshwater systems, which may feature high sediment loads and floating debris. This protocol addresses collecting and analyzing microplastic samples from aquatic environments that contain high suspended solid loads.

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only ...