Name: sediment core sectioning and extraction of pore waters under anoxic conditions
Uploaded: 2016-03-07T13:00:00.000+00:00
Duration: 9 min 21 s
Description: Watch this Scientific Journal Video about Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions at JoVE.com

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. 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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. 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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><tbody><tr><td>Disposable glove bag(s)</td><td>Sigma-Aldrich</td><td>Z106089-1EA</td><td>One per two cores to be processed is usually sufficient.</td></tr><tr><td>N&lt;sub&gt;2&lt;/sub&gt; tank</td><td>Praxair</td><td></td><td>Often gas supply companies can deliver these directly to the field laboratory.</td></tr><tr><td>Nitrogen gas regulator</td><td>VWR</td><td>55850-478</td><td>Or similar</td></tr><tr><td>Several feet of tubing that fits the regulator</td><td>VWR</td><td>89403-862</td><td>Or similar</td></tr><tr><td>Safety equipment to secure the tank</td><td>VWR</td><td>60142-006</td><td></td></tr><tr><td>Adjustable tubing clamp</td><td>VWR</td><td>62849-112</td><td></td></tr><tr><td>Waterproof, good sealing electrical tape</td><td>Scotch</td><td>Super 33+</td><td>Widely available</td></tr><tr><td>2-4 short bungee cords</td><td></td><td></td><td>Widely available</td></tr><tr><td>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><td>Large supply of paper towels and Kimwipes</td><td></td><td></td><td>Widely available</td></tr><tr><td>50 ml centrifuge tubes</td><td>VWR</td><td>21008-951</td><td>Acid cleaned as described in protocol. At least 2/core section needed.</td></tr><tr><td>Several permanent in markers.</td><td></td><td></td><td>Widely available</td></tr><tr><td>Several straight razor blades and box cutters.</td><td></td><td></td><td>Widely available</td></tr><tr><td>Centrifuge</td><td>Beckman-Coulter</td><td>Allegra X-22</td><td>Faster rotor allows greater separation.</td></tr><tr><td>Rotor to accommodate 50 ml tubes</td><td>Beckman-Coulter</td><td>SX-4250</td><td></td></tr><tr><td>50 ml plastic syringes without black rubber tip on the barrel</td><td>VWR</td><td>66064-764&amp;nbsp;</td><td>Acid cleaned as described in protocol. At least 1/core section needed, plus 1 for overlying water.</td></tr><tr><td>Syringe filters compatible with aqueous solutions.</td><td>VWR</td><td>28143-310&amp;nbsp;</td><td>Either 0.45 &amp;mu;m or 0.20&amp;nbsp;&amp;mu;m poresizes may be used. Plan on five filters per core section processed.</td></tr><tr><td>Plastic (disposable) spoons.</td><td></td><td></td><td>Widely available; Acid cleaned as described in protocol.</td></tr><tr><td>Several boxes of disposable gloves.</td><td></td><td></td><td>Widely available</td></tr><tr><td>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><td>Laboratory balance</td><td>VWR</td><td>10205-008</td><td>An available balance will be fine; high precision not required</td></tr><tr><td>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><td>Laboratory notebooks</td><td></td><td></td><td>Water repellent can be useful</td></tr><tr><td>Core liners</td><td>Watermark</td><td>77280</td><td>Available from Forrestry Suppliers</td></tr><tr><td>Core caps</td><td>Ben Meadows</td><td>218105</td><td></td></tr><tr><td>Core slicers</td><td>McMaster Carr</td><td>8707K111</td><td>Cut this into 9 3 x 3 squares</td></tr><tr><td>PVC spacers</td><td>McMaster Carr</td><td>48925K96</td><td>Cut this into short lengths</td></tr><tr><td>PVC couplings</td><td>McMaster Carr</td><td>4880K76</td><td>Approximately 12 needed</td></tr><tr><td>Dowel</td><td></td><td></td><td>Widely available</td></tr><tr><td>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><td>Plywood for core guidance plate and top of lab jack</td><td></td><td></td><td>Widely available</td></tr><tr><td>Lab jack</td><td>VWR</td><td>89260-826</td><td></td></tr><tr><td>Clamps</td><td></td><td></td><td>Widely available</td></tr><tr><td>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

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) loading from lake sediments can account for a substantial portion of the total P load in eutrophic, and some mesotrophic, lakes. Laboratory determination of P release rates from sediment cores is one approach for determining the role of internal P loading and guiding management decisions. Two principal alternatives to experimental determination of sediment P release exist for estimating internal load: in situ measurements of changes in hypolimnetic P over time and P mass balance. The experimental approach using laboratory-based sediment incubations to quantify internal P load is a direct method, making it a valuable tool for lake management and restoration.
Laboratory incubations of sediment cores can help determine the relative importance of internal vs. external P loads, as well as be used to answer a variety of lake management and research questions. We illustrate the use of sediment core incubations to assess the effectiveness of an aluminum sulfate (alum) treatment for reducing sediment P release. Other research questions that can be investigated using this approach include the effects of sediment resuspension and bioturbation on P release.
The approach also has limitations. Assumptions must be made with respect to: extrapolating results from sediment cores to the entire lake; deciding over what time periods to measure nutrient release; and addressing possible core tube artifacts. A comprehensive dissolved oxygen monitoring strategy to assess temporal and spatial redox status in the lake provides greater confidence in annual P loads estimated from sediment core incubations.

Lake eutrophication is a water quality issue worldwide, making the need to identify and control nutrient sources critical. Laboratory determination of phosphorus release rates from sediment cores is a valuable approach for determining the role of internal phosphorus loading and guiding management decisions.

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) ...

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 ...

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled Earth-system processes. Such knowledge is imperative in improving predictions of hydro-biogeochemical cycles, especially under climate change scenarios. We present an experimental method, designed to capture sub-surface heterogeneity of an initially homogeneous soil system. This method is based on destructive sampling of a soil lysimeter designed to simulate a small-scale hillslope. A weighing lysimeter of one cubic meter capacity was divided into sections (voxels) and was excavated layer-by-layer, with sub samples being collected from each voxel. The excavation procedure was aimed at detecting the incipient heterogeneity of the system by focusing on the spatial assessment of hydrological, geochemical, and microbiological properties of the soil. Representative results of a few physicochemical variables tested show the development of heterogeneity. Additional work to test interactions between hydrological, geochemical, and microbiological signatures is planned to interpret the observed patterns. Our study also demonstrates the possibility of carrying out similar excavations in order to observe and quantify different aspects of soil-development under varying environmental conditions and scale.

This study presents an excavation method for investigating subsurface hydrological, geochemical, and microbiological heterogeneity of a soil lysimeter. The lysimeter simulates an artificial hillslope which was initially under homogeneous condition and had been subjected to approximately 5,000 mm of water over eight cycles of irrigation in an 18-month period.

Studying co-evolution of hydrological and biogeochemical processes in the subsurface of natural landscapes can enhance the understanding of coupled ...

Reservoir Condition Pore-scale Imaging of Multiple Fluid Phases Using X-ray Microtomography

X-ray microtomography was used to image, at a resolution of 6.6 &#181;m, the pore-scale arrangement of residual carbon dioxide ganglia in the pore-space of a carbonate rock at pressures and temperatures representative of typical formations used for CO2 storage. Chemical equilibrium between the CO2, brine and rock phases was maintained using a high pressure high temperature reactor, replicating conditions far away from the injection site. Fluid flow was controlled using high pressure high temperature syringe pumps. To maintain representative in-situ conditions within the micro-CT scanner a carbon fiber high pressure micro-CT coreholder was used. Diffusive CO2 exchange across the confining sleeve from the pore-space of the rock to the confining fluid was prevented by surrounding the core with a triple wrap of aluminum foil. Reconstructed brine contrast was modeled using a polychromatic x-ray source, and brine composition was chosen to maximize the three phase contrast between the two fluids and the rock. Flexible flow lines were used to reduce forces on the sample during image acquisition, potentially causing unwanted sample motion, a major shortcoming in previous techniques. An internal thermocouple, placed directly adjacent to the rock core, coupled with an external flexible heating wrap and a PID controller was used to maintain a constant temperature within the flow cell. Substantial amounts of CO2 were trapped, with a residual saturation of 0.203 &#177; 0.013, and the sizes of larger volume ganglia obey power law distributions, consistent with percolation theory.

We present a methodology for the imaging of multiple fluid phases at reservoir conditions by the use of x-ray microtomography. We show some representative results of capillary trapping in a carbonate rock sample.

X-ray microtomography was used to image, at a resolution of 6.6 &#181;m, the pore-scale arrangement of residual carbon dioxide ganglia in the pore-space ...

Engineering

Resurrection of Dormant Daphnia magna: Protocol and Applications

Long-term studies enable the identification of eco-evolutionary processes that occur over extended time periods. In addition, they provide key empirical data that may be used in predictive modelling to forecast evolutionary responses of natural ecosystems to future environmental changes. However, excluding a few exceptional cases, long-term studies are scarce because of logistic difficulties associated with accessing temporal samples. Temporal dynamics are frequently studied in the laboratory or in controlled mesocosm experiments with exceptional studies that reconstruct the evolution of natural populations in the wild.
Here, a standard operating procedure (SOP) is provided to revive or resurrect dormant Daphnia magna, a widespread zooplankton keystone species in aquatic ecosystems, to dramatically advance the state-of-the-art longitudinal data collection in natural systems. The field of Resurrection Ecology was defined in 1999 by Kerfoot and co-workers, even though the first attempts at hatching diapausing zooplankton eggs date back to the late 1980s. Since Kerfoot&#39;s seminal paper, the methodology of resurrecting zooplankton species has been increasingly frequently applied, though propagated among laboratories only via direct knowledge transfer. Here, an SOP is described that provides a step-by-step protocol on the practice of resurrecting dormant Daphnia magna eggs.
Two key studies are provided in which the fitness response of resurrected Daphnia magna populations to warming is measured, capitalizing on the ability to study historical and modern populations in the same settings. Finally, the application of next generation sequencing technologies to revived or still dormant stages is discussed. These technologies provide unprecedented power in dissecting the processes and mechanisms of evolution if applied to populations that have experienced changes in selection pressure over time.

Resurrection of Dormant Daphnia magna: Protocol and Applications

Long-term studies are essential to understanding the process of evolution and the mechanisms of adaptation. Generally, these studies require commitments beyond the life-time of researchers. Here, a powerful method is described that dramatically advances state-of-the-art data collection to generate longitudinal data in natural systems.

Long-term studies enable the identification of eco-evolutionary processes that occur over extended time periods. In addition, they provide key empirical ...

Sediment Core Extrusion Method at Millimeter Resolution Using a Calibrated, Threaded-rod

Aquatic sediment core subsampling is commonly performed at cm or half-cm resolution. Depending on the sedimentation rate and depositional environment, this resolution provides records at the annual to decadal scale, at best. An extrusion method, using a calibrated, threaded-rod is presented here, which allows for millimeter-scale subsampling of aquatic sediment cores of varying diameters. Millimeter scale subsampling allows for sub-annual to monthly analysis of the sedimentary record, an order of magnitude higher than typical sampling schemes. The extruder consists of a 2 m aluminum frame and base, two core tube clamps, a threaded-rod, and a 1 m piston. The sediment core is placed above the piston and clamped to the frame. An acrylic sampling collar is affixed to the upper 5 cm of the core tube and provides a platform from which to extract sub-samples. The piston is rotated around the threaded-rod at calibrated intervals and gently pushes the sediment out the top of the core tube. The sediment is then isolated into the sampling collar and placed into an appropriate sampling vessel (e.g., jar or bag). This method also preserves the unconsolidated samples (i.e., high pore water content) at the surface, providing a consistent sampling volume. This mm scale extrusion method was applied to cores collected in the northern Gulf of Mexico following the Deepwater Horizon submarine oil release. Evidence suggests that it is necessary to sample at the mm scale to fully characterize events that occur on the monthly time-scale for continental slope sediments.

An extrusion method using a calibrated threaded-rod is presented, which allows for mm scale subsampling of aquatic sediment cores. Millimeter-scale sampling is necessary to fully characterize recent event stratigraphy in sediment records.

Aquatic sediment core subsampling is commonly performed at cm or half-cm resolution. Depending on the sedimentation rate and depositional environment, ...

Chemistry

Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors

Denitrification is the primary biogeochemical process removing reactive nitrogen from the biosphere. The quantitative evaluation of this process has become particularly relevant for assessing the anthropogenic-altered global nitrogen cycle and the emission of greenhouse gases (i.e., N2O). Several methods are available for measuring denitrification, but none of them are completely satisfactory. Problems with existing methods include their insufficient sensitivity, and the need to modify the substrate levels or alter the physical configuration of the process using disturbed samples.This work describes a method for estimating sediment denitrification rates that combines coring, acetylene inhibition, and microsensor measurements of the accumulated N2O. The main advantages of this method are a low disturbance of the sediment structure and the collection of a continuous record of N2O accumulation; these enable estimates of reliable denitrification rates with minimum values up to 0.4-1 &#181;mol N2O m-2 h-1. The ability to manipulate key factors is an additional advantage for obtaining experimental insights. The protocol describes procedures for collecting the cores, calibrating the sensors, performing the acetylene inhibition, measuring the N2O accumulation, and calculating the denitrification rate. The method is appropriate for estimating denitrification rates in any aquatic system with retrievable sediment cores. If the N2O concentration is above the detection limit of the sensor, the acetylene inhibition step can be omitted to estimate the N2O emission instead of denitrification. We show how to estimate both actual and potential denitrification rates by increasing nitrate availability as well as the temperature dependence of the process. We illustrate the procedure using mountain lake sediments and discuss the advantages and weaknesses of the technique compared to other methods. This method can be modified for particular purposes; for instance, it can be combined with 15N tracers to assess nitrification and denitrification or field in situ measurements of denitrification rates.

Estimating Sediment Denitrification Rates Using Cores and N2O Microsensors

This method estimates sediment denitrification rates in sediment cores using the acetylene inhibition technique and microsensor measurements of the accumulated N2O. The protocol describes procedures for collecting the cores, calibrating the sensors, performing the acetylene inhibition, measuring the N2O accumulation, and calculating the denitrification rate.

Denitrification is the primary biogeochemical process removing reactive nitrogen from the biosphere. The quantitative evaluation of this process has ...

Isolation of Quartz Grains for Optically Stimulated Luminescence (OSL) Dating of Quaternary Sediments for Paleoenvironmental Research

Optically stimulated luminescence (OSL) dating quantifies the time since mineral grains were deposited and shielded from additional light or heat exposure, which effectively resets the luminescence clock. The systematics of OSL dating is based on the dosimetric properties of common minerals, like quartz and feldspar. The acquired luminescence with exposure to natural ionizing radiation after burial provides a depositional age for many Quaternary sedimentary systems, spanning the past 0.5 Ma. This contribution details the procedures for separating pure quartz grains of a known range of particle sizes to facilitate luminescence analysis with small or single grain aliquots. Specifically, protocols are given for the needed data and interpretations for effective OSL dating of terrestrial sediment cores or sample tubes from exposures. These cores, 5-20 m long in 1.2 m sections, are split lengthwise and crown-cut leaving 80% of core volume undisturbed, which facilitates sampling of light-protected sediment for OSL dating deep within the core. Sediment samples are then subjected to a series of physical separations to obtain a certain grain-size interval (e.g., 150-250 &#181;m). Magnetic minerals are removed in wet and dry states using magnets. A series of chemical digestions starts with soaking in H2O2 to remove organic matter, followed by HCl exposure to remove carbonate minerals, followed by density separation. Subsequently, grains are soaked in HF for 80 min and after in HCl to render solely quartz grains. The mineralogic purity (&#62;99%) of the quartz extract is quantified with grain petrographic assessment and Raman spectroscopy. Repeating this quartz isolation procedure may be necessary with sediment that contains &#60;15% quartz grains. Excitation of the purified quartz grains by LED-derived blue and IR light allows calculations of the fast and IR depletion ratios, which are metrics to assess the dominance of luminescence emissions from quartz.

Isolation of Quartz Grains for Optically Stimulated Luminescence OSL Dating of Quaternary Sediments for Paleoenvironmental Research

This protocol is for the isolation of quartz grains by size for luminescence dating of sediments. Outlined are physical cleansing and chemical digestions by soaking sequentially in H2O2, HCl, HF, and HCl again to isolate quartz grains. The quartz purity is quantified with microscopic assessment, Raman spectroscopy, and IR depletion ratio.

Optically stimulated luminescence (OSL) dating quantifies the time since mineral grains were deposited and shielded from additional light or heat ...

Pore-scale Imaging and Characterization of Hydrocarbon Reservoir Rock Wettability at Subsurface Conditions Using X-ray Microtomography

In situ wettability measurements in hydrocarbon reservoir rocks have only been possible recently. The purpose of this work is to present a protocol to characterize the complex wetting conditions of hydrocarbon reservoir rock using pore-scale three-dimensional X-ray imaging at subsurface conditions. In this work, heterogeneous carbonate reservoir rocks, extracted from a very large producing oil field, have been used to demonstrate the protocol. The rocks are saturated with brine and oil and aged over three weeks at subsurface conditions to replicate the wettability conditions that typically exist in hydrocarbon reservoirs (known as mixed-wettability). After the brine injection, high-resolution three-dimensional images (2 &#181;m/voxel) are acquired and then processed and segmented. To calculate the distribution of the contact angle, which defines the wettability, the following steps are performed. First, fluid-fluid and fluid-rock surfaces are meshed. The surfaces are smoothed to remove voxel artefacts, and in situ contact angles are measured at the three-phase contact line throughout the whole image. The main advantage of this method is its ability to characterize in situ wettability accounting for pore-scale rock properties, such as rock surface roughness, rock chemical composition, and pore size. The in situ wettability is determined rapidly at hundreds of thousands of points. 
The method is limited by the segmentation accuracy and X-ray image resolution. This protocol could be used to characterize the wettability of other complex rocks saturated with different fluids and at different conditions for a variety of applications. For example, it could help in determining the optimal wettability that could yield an extra oil recovery (i.e., designing brine salinity accordingly to obtain higher oil recovery) and to find the most efficient wetting conditions to trap more CO2 in subsurface formations.

This protocol is presented to characterize the complex wetting conditions of an opaque porous medium (hydrocarbon reservoir rock) using three-dimensional images obtained by X-ray microtomography at subsurface conditions.

In situ wettability measurements in hydrocarbon reservoir rocks have only been possible recently. The purpose of this work is to present a protocol to ...

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface in response to disturbances. Deciphering the rapid biogeochemical changes requires in situ, minimally invasive tools with high spatial and temporal sampling resolution. However, the available passive sampling devices are not very useful in many cases either due to their disposable nature or the complexity and extensive workload for sample preparation.
To address this problem, a microdialysis profiler with 33 individual polyethersulfone nanomembrane tubes (semipermeable, &#60;20 nm pore size) integrated into the one-dimensional skeleton (60 mm) was established to iteratively sample the dissolved compounds in porewater across the soil-water interface at a high resolution of 1.8 mm (outer diameter plus one spacing, i.e., 0.1 mm between probes). The sampling mechanism is based on the principle of concentration gradient diffusion. The automatic loading of degassed water allows minimal disturbance to the chemical species across the oxic-anoxic interface.
This paper describes the procedures of device setup and continuous porewater sampling across the soil-water interface on a daily basis. Concentration-depth profiles were selectively measured before (on Day 6) and after (on Day 7) disturbances induced by irrigation. The results showed that concentration-depth profiles were undergoing rapid changes, especially for redox-sensitive elements (i.e., iron and arsenic). These protocols can help investigate the biogeochemical responses across the soil-water interface under various disturbances caused by physical, chemical, and biological factors. The paper thoroughly discusses the advantages and disadvantages of this method for potential use in the environmental sciences.

A microdialysis profiler is described to sample dissolved porewater solutes across an oxic-anoxic soil-water interface in situ with minimal disturbance. This device is designed to capture rapid changes in concentration-depth profiles induced by disturbances at the soil-water interface and beyond.

Biogeochemical processes shift rapidly in both spatial (millimeter scale) and temporal (hour scale to day scale) dimensions at the oxic-anoxic interface ...

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

Summary

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Chapters in this video

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

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

Soil Lysimeter Excavation for Coupled Hydrological, Geochemical, and Microbiological Investigations

Reservoir Condition Pore-scale Imaging of Multiple Fluid Phases Using X-ray Microtomography

Resurrection of Dormant Daphnia magna: Protocol and Applications

Sediment Core Extrusion Method at Millimeter Resolution Using a Calibrated, Threaded-rod

Estimating Sediment Denitrification Rates Using Cores and N₂O Microsensors

Isolation of Quartz Grains for Optically Stimulated Luminescence (OSL) Dating of Quaternary Sediments for Paleoenvironmental Research

Pore-scale Imaging and Characterization of Hydrocarbon Reservoir Rock Wettability at Subsurface Conditions Using X-ray Microtomography

Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers