This work is funded by the National Natural Science Foundation of China (41977320, 41571305) and the Key Programme Special Fund of XJTLU (KSF-A-20).

1. Individual microdialysis sampler preparation
<ol>
	<li>Accurately cut pristine nanomembrane tubes (inner diameter x outer diameter x length: 1.0 mm x 1.7 mm) into a total of 33 short tubes (58 mm in length).</li>
	<li>Accurately cut the PTFE pipe into 66 pipes (180 mm in length) with a ceramic knife. 
	NOTE: Do not use any metal-based knives to avoid any contamination.</li>
	<li>Fully mix the two-part (AB) epoxy adhesive on any clean plastic plate, and let it stand for 30 min until it becomes sticky. Apply the AB epoxy adhesive carefully to the outer surface of the top of the PTFE pipe. Ensure that AB epoxy adhesive only covers the 4 mm length of the tube and that there are no additional adhesive blocking tubes.</li>
	<li>Connect the two PTFE pipes prepared in steps 1.2-1.4 with each nanomembrane tube prepared in step 1.1 by gently screwing the PTFE pipes into the nanomembrane tube. 
	NOTE: Do not allow excess adhesive to accumulate on the joint. Make sure that no adhesive contaminates the nanomembrane tube.</li>
	<li>Repeat step 1.4 to fully assemble all 33 pristine microdialysis samplers.</li>
	<li>Let the samplers assembled in step 1.6 stand overnight to ensure the complete curing and stabilization of the adhesive.</li>
	<li>Enhance the hydrophilicity and clean the microdialysis samplers by soaking them in ethanol (99.5% purity) for 1 h, followed by ultrasonic cleaning (room temperature) with 2% diluted HNO3 and ultrapure water for 15 min each.</li>
	<li>Check the patency and airtightness of the microdialysis sampler by bubbling in water using a 5 mL syringe.</li>
</ol>
2. Assembly of the microdialysis profiler
<ol>
	<li>Use the attached CAD file (Supplemental File 1) to print out the predesigned skeleton using nylon material (Figure 1C-2).</li>
	<li>Hollow out a PVC container (acid-washed) with two parallel slots (5 cm interval) to match the skeleton size. Use the engraving module in the 3D printer for slotting.</li>
	<li>Construct a one-to-many connector by stabilizing epoxy adhesive in the shape of the cap of a 50 mL centrifuge tube. Insert 33 silicon caps (1 cm in length) into the epoxy adhesive before curing, and let stand overnight.</li>
	<li>Take out the one-to-many connector from the tube cap.</li>
	<li>Use a ceramic knife to cut the cured epoxy adhesive so that all the silicon cap ends are unobstructed.</li>
	<li>Rinse the one-to-many connector thoroughly with 2% diluted HNO3 and ultrapure water for 15 min each. Dry the one-to-many connector under ambient conditions.</li>
	<li>Connect a three-way valve to the bottom of the tube to serve as a buffering container.</li>
	<li>Assemble the buffering container by installing a one-to-many connector to a 50 mL centrifuge tube using AB epoxy adhesive.</li>
	<li>Assemble the individual microdialysis samplers prepared in section 1 on the skeleton (step 2.1). In this step, use hot melt adhesive to aid in fixing so that each sampler is parallel to the top/bottom edge of the skeleton.</li>
	<li>Repeat step 2.9 until all the microdialysis samplers (n = 33) are installed on the skeleton.</li>
	<li>Make sure that the 33 samplers on both sides of the skeleton pass through the PVC slots. Seal the gaps at the joints of the skeleton and the slots with AB epoxy adhesive.</li>
	<li>Connect the 33 samplers on one side of the skeleton to a buffering container in an airtight manner via a one-to-many connection valve preinstalled into a 50 mL centrifuge tube (step 2.8)</li>
	<li>Connect a medical infusion bag prefilled with water (18.3 M&#937;) to the buffer container through the three-way valve.</li>
	<li>Use silicon caps to close the 33 samplers on the sampling side. Double-check the patency and airtightness of each microdialysis sampler by turning the three-way valve, allowing water to flow from the medical infusion bag to the sampler. After completing all the checks, close and turn off all the samplers and the valve on the buffering container.</li>
</ol>
3. Soil incubation
<ol>
	<li>Before the incubation of flooded soil, degas the water in the medical infusion bag to remove oxygen. Bubble nitrogen gas overnight in the pathway of the line of high-purity nitrogen gas to the medical infusion bag (Figure 1-C8).</li>
	<li>Use a three-way valve to close the connection between the profiler and the degassed bag.</li>
	<li>Carefully add 450 g of sieved, air-dried soil (particle size &#60; 2 mm) into a PVC container, allowing five microdialysis samplers to remain above the soil surface.</li>
	<li>Use a tissue to cover the soil surface, and then pour ultrapure water (18.3 M&#937;) onto the soil to flood it. Remove the tissue when the soil is fully flooded by 5 cm above the soil surface.</li>
	<li>Purge the system with the preloaded solution immediately once soil incubation has been initialized. Turn on the connection between the anaerobic bag and the dialysis sampler to flush the sampling system. Use 10-fold the total volume of the sampler when purging each sampler with water.</li>
	<li>When finishing the purging of one sampler, cap it using a clean silicon cap.</li>
	<li>Repeat step 3.6 until all the samplers have been purged. At this time, one flooded soil incubation and sampling system are established.</li>
	<li>Adjust the anaerobic bag to the height of the water surface.</li>
	<li>Make sure all the tubes are full of water. If not, remove the cap, and lower the tube top, allowing the water to flow out from the anaerobic bag.</li>
	<li>Close all the caps and valves.</li>
	<li>Turn off the connection between the anaerobic bag and the dialysis sampler during the incubation for 7 days.</li>
</ol>
4. Microdialysis profiler sampling
<ol>
	<li>Before sampling, adjust the water levels in the soil container, the sampling tops, and the anaerobic bag to a similar height to avoid markedly different water potentials. Always maintain this practice over the period of soil incubation.</li>
	<li>Turn on the connection between the anaerobic bag and the buffer container.</li>
	<li>Remove the cap of the first sampler from top to bottom.</li>
	<li>Use a pipette to accurately aspirate 133 &#181;L from the sampler to a vial (0.6 mL) that is preloaded with 133 &#181;L of 2% HNO3 for preservation.</li>
	<li>During the sampling process, observe a slow but uniform flow of water droplets toward the microdialysis sampler in the observation chamber (Figure 1A-9) of the anaerobic bag.</li>
	<li>Close the tube top with a silicon cap. Move to the next sampling tube. 
	NOTE: For the analysis of redox-sensitive elements such as ferrous Fe, a different preservation method such as degassed (10 mM) EDTA solution must be used, and the sampling should be performed under nitrogen purge conditions.</li>
	<li>Repeat step 4.6 until all 33 samples have been collected. Turn off the connection between the anaerobic bag and the buffer container. 
	NOTE: The sampling can generally be finished in 15 min. With the current design, the sampling is carried out on a daily basis to avoid cross-contamination between tubes. Although the solute diffusion along the tube is slow, it would diffuse into the buffer container and contaminate other tubes.</li>
	<li>Immediately after the sampling on Day 6, replenish the flooded water, which will cause a disturbance on the soil surface.</li>
	<li>Calculate the sample volume recovery by weighing the sample vial before and after the porewater sample is transferred.</li>
	<li>Use inductively coupled plasma mass spectrometry (ICP-MS) to measure the total dissolved concentrations of elements in the porewater. 
	NOTE: An external standard curve was used for the concentration quantification, while the internal standard Rh was used for monitoring the ICP-MS operational stability.</li>
</ol>

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The authors declare no conflicts of interest.

Based on previous experiments and practices2, some considerations require special attention during the microdialysis profiler assembly and porewater sampling. First, the nanomembrane tube and the connecting tube should be carefully connected to avoid blockages or leakages at the connection. As the soil is incubated under flooded conditions, the introduction of oxygen will rapidly oxidize and precipitate ferrous iron in the dialysis tubing (Figure 4). For this reason, before assembling the microdialysis profiler, each microdialysis tube must be checked for integrity (no damage), the airtightness of the connections, and the patency of the tubing. Similarly, the connection of the support frame to the side wall of the incubation container needs to be done carefully to avoid leakage. Prior to formal experiments, leakage checks at the various connection locations are always a priority. Second, the perfusate in the anaerobic bag must be adequately deoxygenated. Otherwise, ferrous iron in porewater will react with the oxygen in the perfusate to form insoluble precipitates (Figure 4). This will severely alter the solute speciation and concentration and the diffusion processes toward the nanomembrane tubes. Third, a low sampling frequency (days and weeks) will cause the solute to diffuse into the buffer region. This may contaminate the entire profile sample. To address this problem, three possible solutions can be considered: (1) sampling at a high frequency, such as once a day (however, this may lead to solute depletion near the dialysis sampler when multiple samplings are performed); (2) extending the length of the connecting pipe in the injection area as required; (3) redesigning the sampling pipeline to achieve single control of a single pipeline. These are also directions for the improvement of the device in the future. Fourth, during the sampling process, it must be ensured that the level of the water surface in the anaerobic bag, the flooded soil, and the sampling pipe are approximately at the same height to balance the water pressure. Otherwise, a water potential difference inside and outside the membrane tube will result in a decrease or increase in solute diffusion.
Limitations 
First, since the microdialysis profiler is not commercially available, the method remains time-consuming in terms of the preparation of the device. It took days to prepare a single dialysis tube, including printing the support skeleton, device assembly, and cleaning.&#160;But the subsequent reusable features completely bridge this gap. Second, there are certain limitations in applying the device to non-flooded soil scenarios, which peepers can be used for18. Due to the significant water potential difference between the inside and outside of the membrane tube in dry soil, the preloaded solution experiences diffusion loss; indeed, various sampling volume recoveries in the range of 10%-36% were observed in the preliminary test (detailed data not shown), which creates uncertainty about the results.
Comparison of the method to existing or alternative methods 
The method partially addresses the fact that the existing passive samplers cannot sample repeatedly and minimizes the workload of sample preparation, especially for anoxic porewater sampling and preservation2. The instant changes in concentration and speciation of dialyzed solutes can sensitively reflect the response of the oxic-anoxic interface to any environmental disturbances. Theoretically, sampling at a frequency of minutes, hours, or days allows for the capture of the rapidly changing processes at the interface. For passive samplers that need to be in deployment for days, some hot moments&#160;and hotspots can be missed6,19.
Importance and potential applications in the environmental sciences 
This approach could advance biogeochemistry studies at oxic-anoxic interfaces, for instance, to find hot moments and hotspots of biogeochemical processes under specific Eh-pH conditions. The redox process is the basic process of life activities1. Microorganisms, especially, require optimal living environment conditions and are very sensitive to environmental disturbances1. This results in a highly dynamic development of microbial communities and biogeochemical processes in heterogeneous environments20. Direct sampling, without considering the high heterogeneity, tends to obtain a mixed sample from various environmental conditions. This causes mismatches between the measured chemical information and key microorganisms20. Within a few centimeters of the surface layer of soil or sediment in a typical flooded paddy field, there are steep redox gradients, as well as various physical, chemical21, and biological gradients1. Technology must be able to capture millimeter-scale biogeochemical signals; otherwise, data that do not match the actual scale may lead to ambiguous conclusions. The microdialysis profiler is capable of monitoring millimeter-scale biochemical signals at the soil-water interface in days or hours with minimal disturbance. In this study, the spatiotemporal dynamics of different elements over a 48 h period were observed, possibly related to the disturbance of water replenishment. Therefore, a broader application of the microdialysis profiler may help to understand how disturbances affect key biogeochemical processes in a changing world.

An oxic-anoxic interface is one general feature in the biosphere that is vital for the biogeochemical cycle1. This interface is highly heterogeneous, with the spatial range extending from millimeters in the sediment/soil-water interface1,2 to thousands of meters in the oceanic anoxic zone3,4. This interface is an ideal habitat for studying the complexity of elemental biogeochemistry.
Soil-water interfaces have a typical oxic-anoxic gradient feature within centimeters and are easily established in mesocosm experiments. Starting from the consumption of molecular oxygen from surface water, the stratified functional microbial communities drive the development of various gradients, such as O2, pH, and Eh gradients, at the millimeter scale1. Biogeochemical cycling at the oxic-anoxic interface is sensitive to various disturbances in nature5,6. In the case of sediments and paddy fields, the input of fresh organic matter such as litter and straw, periodic flooding and drainage, temperature fluctuations and extremes, and bioturbation can cause changes in the biogeochemical cycle at the oxic-anoxic interface, likely resulting in lasting impacts, such as greenhouse gas emissions, eutrophication, and contamination at a given location. Therefore, the oxic-anoxic gradient at the soil-water interface provides a window for the study of global, large-scale, biogeochemical cycles. The spatiotemporal sampling and analysis of dissolved substances along the soil-water interface in high resolution have always been of interest; however, there has been limited progress in the methodology.
Circumventing the drawbacks of destructive porewater extraction, non-destructive passive sampling is increasingly used to avoid changes in porewater chemistry and address the complexity of sample preparation7. Several devices that can perform high-precision, in situ sampling (from the micrometer to centimeter scale) have been widely used, including in situ dialysis samplers (known as peepers)8, diffusive equilibration in thin films (DET)9, and diffusive gradient in thin films (DGT)10. Dissolved substances are passively sampled via the mechanism of diffusion and adsorption processes. Although they have proven useful in describing oxic-anoxic chemical profiles, they are still single-use, which limits their broader application.
Recently, the microdialysis technique has emerged as a sensitive tool that can be used to monitor soluble compound dynamics in soil on temporal scales of minutes to days11,12,13,14. For a typical scenario using microdialysis in medical and environmental sciences, a miniature, concentric-type probe consisting of a semipermeable tubular membrane (i.e., a microdialyzer) is used to probe the interstitial fluid or soil solutions to prevent significant disturbances on, metabolic processes&#160;and chemical speciation15,16. One of the greatest inherent advantages of microdialysis is the in situ capture of time-dependent concentration changes in soil or biological tissues15,16.
Based on the microdialysis concept, we developed a more easy-to-use microdialysis profiler, previously called the integrated porewater injection&#160;(IPI) profiler, that can perform continuous equilibrium dialysis of porewater solutes based on the principle of concentration gradient diffusion2. The microdialysis device uses hollow nanomembrane tubes for active preloading of the perfusate and passive diffusion of the dissolved solutes, which is different from the bulk porewater diffusion used in peepers, pressure filters such as the Rhizon sampler, and accumulation-based DGT. The device has been tested and validated in the temporal and spatial sampling of both cationic and anionic elements in both highland and flooded soils (Figure 1A-1)13,15,16. Simple pump in-and-out microdialysis minimizes the number of steps in sample preparation2,15.
We fabricated a microdialysis profiler by integrating a set of samplers onto a one-dimensional support skeleton, and this profiler achieved high-resolution sampling at the soil-water interface and rhizosphere2,15,17. In this study, considerable modifications of the sampling device and sampling method were made to allow the collection of 33 pore water samples at the soil-water interface (60 mm vertical depth) with minimal disturbance for downstream elemental analysis. The whole sampling procedure takes ~15 min. Since the microdialysis profiler is new to the community of environmental science, we present details of the device components and sampling procedures to indicate the potential of microdialysis in monitoring the changes in chemical signals at the soil-water interface.
Description of the microdialysis profiler 
The microdialysis profiler device, with proper modifications of the previous design2, is shown in Figure 1. The effective pore size of the nanomembrane (Figure 1C-1) is estimated to be only several nanometers to prevent the diffusion of large molecules and microbial cells. A previous test suggested that a 6 month flooded incubation did not result in any iron deposits on either the inside or the outside of the tube surface15. A curved, hollow skeleton was designed (Figure 1C-2) and 3D-printed using a stable nylon material. A total of 33 nanomembrane tubes (polyethersulfone; surface pore size: 0-20 nm; inner diameter x outer diameter x effective sampling length: 1.0 mm x 1.7 mm x 54 mm; theoretical volume: 42.4 &#181;L) connected with matched polytetrafluoroethylene (PTFE) pipes (length: 18 cm x 2 cm diameter Figure 1C-1) were installed on the skeleton and across one side of a PVC container (Figure 1B). For this device, the sampling component (Figure 1B-1) is 2 cm away from the side wall of the PVC container. For the injection side (Figure 1B-4), all tubes were connected to a one-to-many connector, which was fixed into a buffering container in an airtight manner (Figure 1B-7). A medical infusion bag (Figure 1B-11) was used to connect with the buffering container by a three-way valve. The airtightness of the system was carefully examined in water before further experimental operations. The preloaded water (18.2 M&#937;, 500 mL) in the medical infusion bag is always oxygen-free (Figure 1C-8). Detailed device setup and porewater sampling are described as follows.

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	<tbody>
		<tr>
			<td>3D Printer</td>
			<td>Snapmaker, United States</td>
			<td>Snapmaker 2.0</td>
			<td>&#160;Model: A250</td>
		</tr>
		<tr>
			<td>3M DP190 Scotch-Weld Gray</td>
			<td>&#160;3M United States</td>
			<td>489-483</td>
			<td>Gray</td>
		</tr>
		<tr>
			<td>Centrifuge tube</td>
			<td>Titan, China</td>
			<td>SWLX-JZ050-ZX</td>
			<td>50 mL, Sterilized DNASE/RNASE/Protease/Pyrogen Free</td>
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			<td>Ceramic knife</td>
			<td>R felngli, China</td>
			<td>N.A.</td>
			<td>General</td>
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			<td>EDTA FREE ACID</td>
			<td>Sigma-Aldrich</td>
			<td>CAS 60-00-4</td>
			<td>Sigma-Aldrich#EDS-1KG</td>
		</tr>
		<tr>
			<td>Ethanol</td>
			<td>Adamas</td>
			<td>CAS 64-17-5</td>
			<td>Water &#8804; 50 ppm (by K.F.), 99.5%, SafeDry, with molecular sieves, Safeseal</td>
		</tr>
		<tr>
			<td>Hot melt adhesive&#160;</td>
			<td>Magic Dragon, China</td>
			<td>N.A.</td>
			<td>JTWJRRJB001</td>
		</tr>
		<tr>
			<td>Inductively Coupled Plasma Mass Spectrometry&#160;</td>
			<td>PerkinElmer, Inc., Shelton, CT USA</td>
			<td>N.A.</td>
			<td>Model: NexION 350X</td>
		</tr>
		<tr>
			<td>Medical Infusion Bag&#160;</td>
			<td>Hunan Kanglilai Medical Equipment Co., Ltd</td>
			<td>N.A.</td>
			<td>250 Ml, Sterlized</td>
		</tr>
		<tr>
			<td>Milli-Q water system</td>
			<td>Mingche, Inc., China</td>
			<td>N.A.</td>
			<td>18.3 M&#937;, water purification system model: 24UV</td>
		</tr>
		<tr>
			<td>Nanomembrane Tube (polyethersulfone)</td>
			<td>Motimo Membrane Technology Co., Ltd., Tianjin, China</td>
			<td>N.A.</td>
			<td>Polyethersulfone, inner diameter 1 mm, poresize &#60;20 nm, pretreated with ethanol (99.5%)</td>
		</tr>
		<tr>
			<td>Nitrogen gas</td>
			<td>Suzhou Gas, Chuina</td>
			<td>N.A.</td>
			<td>High puriety</td>
		</tr>
		<tr>
			<td>Nitrotic acid (Concentrated)</td>
			<td>Adamas</td>
			<td>CAS 7697-37-2</td>
			<td>69%,Single Metal &#60; 50 ppt, PFA Bottle</td>
		</tr>
		<tr>
			<td>Nylon Fiber</td>
			<td>Soumiety</td>
			<td>10052076600273</td>
			<td>For 3D-printing</td>
		</tr>
		<tr>
			<td>Pipette&#160;</td>
			<td>Bond A3 Pipette</td>
			<td>N.A.</td>
			<td>200 &#956;L</td>
		</tr>
		<tr>
			<td>Pipette Tip</td>
			<td>Titan</td>
			<td>T2-H-T0200</td>
			<td>200 &#956;L, 300 &#956;L Tip Box Non-sterile|200 &#956;L|Titan</td>
		</tr>
		<tr>
			<td>Polytetrafluoroethylene Tube</td>
			<td>ROHS, China</td>
			<td>CJ-TTL</td>
			<td>Out diameter 1 mm</td>
		</tr>
		<tr>
			<td>Sample vial</td>
			<td>Titan, China</td>
			<td>EP0060-B-N</td>
			<td>0.6 mL, Sterilized DNASE/RNASE/Protease/Pyrogen Free</td>
		</tr>
		<tr>
			<td>Silicon cap</td>
			<td>Fuchenxiangsu, China</td>
			<td>N.A.</td>
			<td>Inner diameter 1 mm, length 1 cm</td>
		</tr>
		<tr>
			<td>Sonicator</td>
			<td>Elma</td>
			<td>N.A.</td>
			<td>model:E120H</td>
		</tr>
		<tr>
			<td>Square PVC water pipe</td>
			<td>Taobao.com</td>
			<td>N.A.</td>
			<td>hight x width, 12 cm x 15 cm</td>
		</tr>
		<tr>
			<td>Three-way valve for infusion</td>
			<td>OEM, China</td>
			<td>N.A.</td>
			<td>Medical level; Valve body: PC material; valve core: PE material; screw cap: ABS material</td>
		</tr>
	</tbody>
</table>

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.

Following this protocol, a microdialysis profiler system was established, as described in Figure 1. Soil incubation was performed under flooding conditions (24 &#176;C, uncovered from light). Samples on Day 6 and Day 7 were selectively measured to indicate potential disturbance on the soil surface due to the practice of replenishing the flooded water.
During each sampling, a consistent number of water droplets in the observation chamber flowing toward the microdialysis sampler was observed, indicating that the transferred sample solution was continuously replenished by the solution in the anaerobic bag. As shown in Figure 2, the recovery percentage of the sample volume averaged 101.4% &#177; 0.9% and ranged from 100.2% to 103.6%. A slightly higher recovery of the sample volume might indicate that there was a water level difference between the anaerobic bag and the top of the sampling tube.
Using the samples across the soil-water interface collected on Day 6 and Day 7, the total dissolved concentrations of iron (Fe), manganese (Mn), arsenic (As), cadmium (Cd), copper (Cu), lead (Pb), nickel (Ni), and zinc (Zn) in the porewater were determined (Figure 3). The concentration-depth profiles varied greatly depending on the elemental type and before and after the practice of replenishing the flooded water. Although we did not perform replications here since this study used a gradient-based experimental design, our previous study demonstrated good replications of changes in depth-dependent chemical signals18.
On Day 6, the dissolved concentrations of Mn, Fe, and As increased along with the soil depth, whereas those of Cu and Pb decreased with increasing soil depth. The results are consistent with the general principles and observations in soil-water interfaces; specifically, a more reduced environment in deeper soil would cause an enhanced reductive release of Mn15, Fe, and As while inhibiting the release of cationic metals due to the formation of less soluble minerals. However, for Cd, Ni, and Zn, the concentration-depth profiles indicated a different pattern, since the dissolved concentrations had an increasing trend from a depth of around &#8722;20 mm to deeper locations.
Compared to the concentration-depth profiles of Fe (4.95 mg&#183;L&#8722;1) and As (3.3 &#181;g&#183;L&#8722;1) at the depth of &#8722;12 mm on Day 6, the concentrations of Fe (1.46 mg&#183;L&#8722;1) and As (0.8 &#181;g&#183;L&#8722;1) were significantly lower on Day 7; however, the Fe and As concentrations were significantly higher (depth-dependent slope, p &#60; 0.001) from the depths of &#8722;18 mm to &#8722;50 mm. For most elements determined, except Mn, the dissolved concentrations in the surface water and the even surface soil at the depth of &#8722;15 mm were significantly lower, to varying degrees, after aerobic water replenishment. It was noted that there was a concentration peak for Pb at the depth of approximately &#8722;10 mm on Day 7, showing a contrasting pattern to that observed on Day 6. These inconsistent results are likely caused by the disturbance of water replenishment and the temporal evolution of biogeochemistry across the soil-water interface. In either case, the microdialysis profiler indicated its great potential to monitor the temporospatial changes in chemical profiles across the soil-water interface.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/64358/64358fig01.jpg" /> 
Figure 1: Microdialysis profiler setup for monitoring chemical dynamics at soil-water interfaces to the soil depth of 50 mm. (A) For a profiler in use at 50 mm depth, seealso Supplementary Figure S1. The main components include (B1,C1) 33 microdialysis samplers (B2,C2) installed on a 3D-printed skeleton, which is further installed on a (B3) incubation container (a 50 mL sample tube), (B4,B7,C4) a one-to-many buffering container, (B9-B12) a medical infusion bag used as the supplier of degassed water, and an (C5) offline sampling pipette. (B5) The sampling locations of all 33 samplers are aligned to the same height with (B6) a plastic strip. Deoxygenated water is prepared by (C8) nitrogen bubbling in a reverse direction to the water supply. <a href="https://www.jove.com/files/ftp_upload/64358/64358fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/64358/64358fig02.jpg" /> 
Figure 2: Sampling volume recovery using H2O as the perfusate. The error bars denote the standard deviation of two independent profiler samplings. <a href="https://www.jove.com/files/ftp_upload/64358/64358fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/64358/64358fig03.jpg" /> 
Figure 3: Concentration-depth profiles. (A) Manganese, (B) iron, (C) arsenic, (D) cadmium, (E) copper, (F) lead, (G) nickel, and (H) zinc measured on Day 6 and Day 7. The negative tick labels on the Y-axis indicate the depths below the water-soil boundary. <a href="https://www.jove.com/files/ftp_upload/64358/64358fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/64358/64358fig04.jpg" /> 
Figure 4: Failure case of leakage resulting in iron precipitation inside the samplers. <a href="https://www.jove.com/files/ftp_upload/64358/64358fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary File 1: Computer-assisted design file for a printout of the predesigned skeleton. <a href="https://www.jove.com/files/ftp_upload/64358/3D printing file.zip" target="_blank">Please click here to download this File.</a>
Supplementary Figure S1: The profiler in use. (A) On flooded soil. (B-E) Photos of top and side views and connection details are presented separately. (E) Three-way valves are used to connect the buffer container and the medical infusion bag. <a href="https://www.jove.com/files/ftp_upload/64358/64358_Supplemental Fig S1.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Dissolved Solute Sampling Across an Oxic-Anoxic Soil-Water Interface Using Microdialysis Profilers at JoVE.com

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

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 ...

dissolved-solute-sampling-across-an-oxic-anoxic-soil-water-interface

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1.2K Views.  University of Liverpool. 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.

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

Soil Sampling and Isolation of Entomopathogenic Nematodes (Steinernematidae, Heterorhabditidae)

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to two families: Steinernematidae and Heterorhabditidae. Until now, more than 70 species have been described in the Steinernematidae and there are about 20 species in the Heterorhabditidae. The nematodes have a mutualistic partnership with Enterobacteriaceae bacteria and together they act as a potent insecticidal complex that kills a wide range of insect species.
Herein, we focus on the most common techniques considered for collecting EPN from soil. The second part of this presentation focuses on the insect-baiting technique, a widely used approach for the isolation of EPN from soil samples, and the modified White trap technique which is used for the recovery of these nematodes from infected insects. These methods and techniques are key steps for the successful establishment of EPN cultures in the laboratory and also form the basis for other bioassays that consider these nematodes as model organisms for research in other biological disciplines. The techniques shown in this presentation correspond to those performed and/or designed by members of S. P. Stock laboratory as well as those described by various authors.

Soil Sampling and Isolation of Entomopathogenic Nematodes Steinernematidae, Heterorhabditidae

Entomopathogenic nematodes are soil-inhabiting roundworms that parasitize a wide range of insects. We demonstrate sampling methods used for the isolation of these nematodes from the soil using two techniques: the insect baiting and the modified White trap, for the recovery of nematodes from soil samples and infected insect cadavers, respectively.

Entomopathogenic nematodes (a.k.a. EPN) represent a group of soil-inhabiting nematodes that parasitize a wide range of insects. These nematodes belong to ...

Spotting Cheetahs: Identifying Individuals by Their Footprints

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges widely over sub-Saharan Africa and in parts of the Middle East. Cheetah conservationists face two major challenges, conflict with landowners over the killing of domestic livestock, and concern over range contraction. Understanding of the latter remains particularly poor2. Namibia is believed to support the largest number of cheetahs of any range country, around 30%, but estimates range from 2,9053 to 13,5204. The disparity is likely a result of the different techniques used in monitoring.
Current techniques, including invasive tagging with VHF or satellite/GPS collars, can be costly and unreliable. The footprint identification technique5 is a new tool accessible to both field scientists and also citizens with smartphones, who could potentially augment data collection. The footprint identification technique analyzes digital images of footprints captured according to a standardized protocol. Images are optimized and measured in data visualization software. Measurements of distances, angles, and areas of the footprint images are analyzed using a robust cross-validated pairwise discriminant analysis based on a customized model. The final output is in the form of a Ward's cluster dendrogram. A user-friendly graphic user interface (GUI) allows the user immediate access and clear interpretation of classification results.
The footprint identification technique algorithms are species specific because each species has a unique anatomy. The technique runs in a data visualization software, using its own scripting language (jsl) that can be customized for the footprint anatomy of any species. An initial classification algorithm is built from a training database of footprints from that species, collected from individuals of known identity. An algorithm derived from a cheetah of known identity is then able to classify free-ranging cheetahs of unknown identity. The footprint identification technique predicts individual cheetah identity with an accuracy of &gt;90%.

The cheetah (Acinonyx jubatus) is an iconic, endangered species, but conservation efforts are challenged by habitat shrinkage and conflict with commercial farmers. The footprint identification technique, a robust, accurate and cost-effective image classification system, is a new approach to monitoring cheetahs.

The cheetah (Acinonyx jubatus) is Africa's most endangered large felid and listed as Vulnerable with a declining population trend by the IUCN1. It ranges ...

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 ...

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and detritus, and excreting particulate and dissolved compounds, they serve as important agents for benthic-pelagic coupling. Accurately measuring the compounds removed and excreted by suspension feeders (such as sponges, ascidians, polychaetes, bivalves) is crucial for the study of their physiology, metabolism, and feeding ecology, and is fundamental to determine the ecological relevance of the nutrient fluxes mediated by these organisms. However, the assessment of the rate by which suspension feeders process particulate and dissolved compounds in nature is restricted by the limitations of the currently available methodologies. Our goal was to develop a simple, reliable, and non-intrusive method that would allow clean and controlled water sampling from a specific point, such as the excurrent aperture of benthic suspension feeders, in situ. Our method allows simultaneous sampling of inhaled and exhaled water of the studied organism by using minute tubes installed on a custom-built manipulator device and carefully positioned inside the exhalant orifice of the sampled organism. Piercing a septum on the collecting vessel with a syringe needle attached to the distal end of each tube allows the external pressure to slowly force the sampled water into the vessel through the sampling tube. The slow and controlled sampling rate allows integrating the inherent patchiness in the water while ensuring contamination free sampling. We provide recommendations for the most suitable filtering devices, collection vessel, and storing procedures for the analyses of different particulate and dissolved compounds. The VacuSIP system offers a reliable method for the quantification of undisturbed suspension feeder metabolism in natural conditions that is cheap and easy to learn and apply to assess the physiology and functional role of filter feeders in different ecosystems.

VacuSIP, an Improved InEx Method for In Situ Measurement of Particulate and Dissolved Compounds Processed by Active Suspension Feeders

We introduce the VacuSIP, a simple, non-intrusive, and reliable method for clean and accurate point sampling of water. The system was developed and evaluated for the simultaneous collection of the water inhaled and exhaled by benthic suspension feeders in situ, to cleanly measure removal and excretion of particulate and dissolved compounds.

Benthic suspension feeders play essential roles in the functioning of marine ecosystems. By filtering large volumes of water, removing plankton and ...

A Simple Approach to Manipulate Dissolved Oxygen for Animal Behavior Observations

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal behavior questions. The protocol described here provides a simple, reproducible, and controlled method to manipulate DO to study behavioral response in aquatic organisms resulting from hypoxic and anoxic conditions. While performing degasification of water with nitrogen is commonly used in laboratory settings, no explicit method for ecological (aquatic) application exists in the literature, and this protocol is the first to describe a protocol to degasify water to observe organismal response. This technique and protocol were developed for direct application for aquatic macroinvertebrates; however, small fish, amphibians, and other aquatic vertebrates could be easily substituted. It allows for easy manipulation of DO levels ranging from 2 mg/L to 11 mg/L with stability for up to a 5 min animal-observation period. Beyond a 5 min observation period water temperatures began to rise, and at 10 min DO levels became too unstable to maintain. The protocol is scalable to the study organism, reproducible, and reliable, allowing for rapid implementation into introductory teaching labs and high-level research applications. The expected results of this technique should relate dissolved oxygen changes to behavioral responses of organisms.

This article describes a simple and reproducible protocol to manipulate dissolved oxygen conditions in a laboratory setting for animal behavior studies. This protocol may be used in both teaching and research laboratory settings to evaluate organismal response of macroinvertebrates, fishes, or amphibians to changes in dissolved oxygen concentration.

The ability to manipulate dissolved oxygen (DO) in a laboratory setting has significant application to investigate a number of ecological and organismal ...

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 ...

Understanding Dissolved Organic Matter Biogeochemistry Through In Situ Nutrient Manipulations in Stream Ecosystems

Dissolved organic matter (DOM) is a highly diverse mixture of molecules providing one of the largest sources of energy and nutrients to stream ecosystems. Yet the in situ study of DOM is difficult as the molecular complexity of the DOM pool cannot be easily reproduced for experimental purposes. Nutrient additions to streams however, have been shown to repeatedly alter the in situ and ambient DOM pool. Here we demonstrate an easily replicable field-based method for manipulating the ambient pool of DOM at the ecosystem scale. During nutrient pulse experiments changes in the concentration of both dissolved organic carbon and dissolved organic nitrogen can be examined across a wide-range of nutrient concentrations. This method allows researchers to examine the controls on the DOM pool and make inferences regarding the role and function that certain fractions of the DOM pool play within ecosystems. We advocate the use of this method as a technique to help develop a deeper understanding of DOM biogeochemistry and how it interacts with nutrients. With further development this method may help elucidate the dynamics of DOM in other ecosystems.

Understanding Dissolved Organic Matter Biogeochemistry Through In Situ Nutrient Manipulations in Stream Ecosystems

Dissolved organic matter provides an important source of energy and nutrients to stream ecosystems. Here we demonstrate a field-based method to manipulate the ambient pool of dissolved organic matter in situ through easily replicable nutrient pulses.

Dissolved organic matter (DOM) is a highly diverse mixture of molecules providing one of the largest sources of energy and nutrients to stream ecosystems. ...

A Lipid Extraction and Analysis Method for Characterizing Soil Microbes in Experiments with Many Samples

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial communities, a relatively precise but effort-intensive technique to assay microbial community composition is phospholipid fatty acid (PLFA) analysis. PLFA was developed to analyze phospholipid biomarkers, which can be used as indicators of microbial biomass and the composition of broad functional groups of fungi and bacteria. It has commonly been used to compare soils under alternative plant communities, ecology, and management regimes. The PLFA method has been shown to be sensitive to detecting shifts in microbial community composition.
An alternative method, fatty acid methyl ester extraction and analysis (MIDI-FA) was developed for rapid extraction of total lipids, without separation of the phospholipid fraction, from pure cultures as a microbial identification technique. This method is rapid but is less suited for soil samples because it lacks an initial step separating soil particles and begins instead with a saponification reaction that likely produces artifacts from the background organic matter in the soil. 
This article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples. The method combines the accuracy achieved through PLFA profiling by extracting and concentrating soil lipids as a first step, and a reduction in effort by saponifying the organic material extracted and processing with the MIDI-FA method as a second step.

The article describes a method that increases throughput while balancing effort and accuracy for extraction of lipids from the cell membranes of microorganisms for use in characterizing both total lipids and the relative abundance of indicator lipids to determine soil microbial community structure in studies with many samples.

Microbial communities are important drivers and regulators of ecosystem processes. To understand how management of ecosystems may affect microbial ...

Sampling Soils in a Heterogeneous Research Plot

Soils are highly heterogeneous. In general, the number of soil samples required for soil research has always been determined arbitrarily and the associated accuracy is unknown. Here, we present a detailed protocol for efficient and clustered soil sampling in a research plot and, relying on a pilot sampling using this design, for demonstrating soil spatial heterogeneity and informing reasonable sample sizes and associated accuracy for future study. The protocol mainly comprises four steps: sampling design, field collection, soil analysis, and geostatistical analysis. The step-by-step procedure is modified according to former publications. Two examples will be presented to demonstrate contrasting spatial distributions of soil organic carbon (SOC) and soil microbial biomass carbon (MBC) under different management practices. In addition, we present a strategy to determine the sample size requirement (SSR) given a certain level of accuracy based on the plot-level coefficient of variation (CV). The field sampling protocol and the quantitative determination of the sample size will assist researchers in seeking feasible sampling strategies to meet research needs and resources' availability.

The traditional soil-sampling procedure determines the number of soil samples arbitrarily. Here, we provide a simple yet efficient clustered soil-sampling design to demonstrate soil spatial heterogeneity and quantitatively determine the number of soil samples required and the associated sampling accuracy.

Soils are highly heterogeneous. In general, the number of soil samples required for soil research has always been determined arbitrarily and the ...

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

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

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