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>

<ol>
	<li>Brune, A., Frenzel, P., Cypionka, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Life+at+the+oxic-anoxic+interface:+Microbial+activities+and+adaptations.">Life at the oxic-anoxic interface: Microbial activities and adaptations.</a> FEMS Microbiology Reviews. 24 (5), 691-710 (2000).</li><li>Yuan, Z. -. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Tracing+the+dynamic+changes+of+element+profiles+by+novel+soil+porewater+samplers+with+ultralow+disturbance+to+soil-water+interface.">Tracing the dynamic changes of element profiles by novel soil porewater samplers with ultralow disturbance to soil-water interface.</a> Environmental Science &amp; Technology. 53 (9), 5124-5132 (2019).</li><li>Henkel, S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Diagenetic+barium+cycling+in+Black+Sea+sediments+-+A+case+study+for+anoxic+marine+environments.">Diagenetic barium cycling in Black Sea sediments - A case study for anoxic marine environments.</a> Geochimica et Cosmochimica Acta. 88, 88-105 (2012).</li><li>Zhong, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Novel+insights+into+the+Thaumarchaeota+in+the+deepest+oceans:+Their+metabolism+and+potential+adaptation+mechanisms.">Novel insights into the Thaumarchaeota in the deepest oceans: Their metabolism and potential adaptation mechanisms.</a> Microbiome. 8 (1), 78 (2020).</li><li>Lueder, U., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Influence+of+physical+perturbation+on+Fe(II)+supply+in+coastal+marine+sediments.">Influence of physical perturbation on Fe(II) supply in coastal marine sediments.</a> Environmental Science &amp; Technology. 54 (6), 3209-3218 (2020).</li><li>Sharma, N., Wang, Z., Catalano, J. G., Giammar, D. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+responses+of+trace+metal+bioaccessibility+to+fluctuating+redox+conditions+in+wetland+soils+and+stream+sediments.">Dynamic responses of trace metal bioaccessibility to fluctuating redox conditions in wetland soils and stream sediments.</a> ACS Earth and Space Chemistry. 6 (5), 1331-1344 (2022).</li><li>Vrana, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Passive+sampling+techniques+for+monitoring+pollutants+in+water.">Passive sampling techniques for monitoring pollutants in water.</a> TrAC Trends in Analytical Chemistry. 24 (10), 845-868 (2005).</li><li>VanOploo, P., White, I., Macdonald, B. C. T., Ford, P., Melville, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+use+of+peepers+to+sample+pore+water+in+acid+sulphate+soils.">The use of peepers to sample pore water in acid sulphate soils.</a> European Journal of Soil Science. 59 (4), 762-770 (2008).</li><li>Harper, M. P., Davison, W., Tych, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporal,+spatial,+and+resolution+constraints+for+in+situ+sampling+devices+using+diffusional+equilibration:+Dialysis+and+DET.">Temporal, spatial, and resolution constraints for in situ sampling devices using diffusional equilibration: Dialysis and DET.</a> Environmental Science &amp; Technology. 31 (11), 3110-3119 (1997).</li><li>Harper, M. P., Davison, W., Tych, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Estimation+of+pore+water+concentrations+from+DGT+profiles:+a+modelling+approach.">Estimation of pore water concentrations from DGT profiles: a modelling approach.</a> Aquatic Geochemistry. 5 (4), 337-355 (1999).</li><li>Gao, S., DeLuca, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+microdialysis+to+assess+short-term+soil+soluble+N+dynamics+with+biochar+additions.">Use of microdialysis to assess short-term soil soluble N dynamics with biochar additions.</a> Soil Biology and Biochemistry. 136, 107512 (2019).</li><li>Buckley, S., Brackin, R., J&#228;mtg&#229;rd, S., N&#228;sholm, T., Schmidt, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microdialysis+in+soil+environments:+Current+practice+and+future+perspectives.">Microdialysis in soil environments: Current practice and future perspectives.</a> Soil Biology and Biochemistry. 143, 107743 (2020).</li><li>Mir&#243;, M., Jimoh, M., Frenzel, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+dynamic+approach+for+automatic+microsampling+and+continuous+monitoring+of+metal+ion+release+from+soils+exploiting+a+dedicated+flow-through+microdialyser.">A novel dynamic approach for automatic microsampling and continuous monitoring of metal ion release from soils exploiting a dedicated flow-through microdialyser.</a> Analytical and Bioanalytical Chemistry. 382 (2), 396-404 (2005).</li><li>Maddala, S., Savin, M. C., Stenken, J. A., Wood, L. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Nitrogen+dynamics:+Quantifying+and+differentiating+fluxes+in+a+riparian+wetland+soil.">Nitrogen dynamics: Quantifying and differentiating fluxes in a riparian wetland soil.</a> ACS Earth and Space Chemistry. 5 (5), 1254-1264 (2021).</li><li>Yuan, Z. -. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Simultaneous+measurement+of+aqueous+redox-sensitive+elements+and+their+species+across+the+soil-water+interface.">Simultaneous measurement of aqueous redox-sensitive elements and their species across the soil-water interface.</a> Journal of Environmental Sciences. 102, 1-10 (2021).</li><li>Hamilton, E. M., Young, S. D., Bailey, E. H., Humphrey, O. S., Watts, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Online+microdialysis-high-performance+liquid+chromatography-inductively+coupled+plasma+mass+spectrometry+(MD-HPLC-ICP-MS)+as+a+novel+tool+for+sampling+hexavalent+chromium+in+soil+solution.">Online microdialysis-high-performance liquid chromatography-inductively coupled plasma mass spectrometry (MD-HPLC-ICP-MS) as a novel tool for sampling hexavalent chromium in soil solution.</a> Environmental Science &amp; Technology. 55 (4), 2422-2429 (2021).</li><li>Yuan, Z. -. F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Distinct+and+dynamic+distributions+of+multiple+elements+and+their+species+in+the+rice+rhizosphere.">Distinct and dynamic distributions of multiple elements and their species in the rice rhizosphere.</a> Plant and Soil. 471 (1), 47-60 (2022).</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 Trends in Analytical Chemistry. 14 (6), 250-256 (1995).</li><li>Wey, H., Hunkeler, D., Bischoff, W. -. A., B&#252;nemann, E. K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Field-scale+monitoring+of+nitrate+leaching+in+agriculture:+assessment+of+three+methods.">Field-scale monitoring of nitrate leaching in agriculture: assessment of three methods.</a> Environmental Monitoring and Assessment. 194 (1), (2021).</li><li>Cai, Y. -. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Microbial+community+structure+is+stratified+at+the+millimeter-scale+across+the+soil-water+interface.">Microbial community structure is stratified at the millimeter-scale across the soil-water interface.</a> ISME Communications. 2 (1), 53 (2022).</li><li>Jones, M. E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Manganese-driven+carbon+oxidation+at+oxic-anoxic+interfaces.">Manganese-driven carbon oxidation at oxic-anoxic interfaces.</a> Environmental Science &amp; Technology. 52 (21), 12349-12357 (2018).</li></ol>

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.

<table><tbody><tr><td>3D Printer</td><td>Snapmaker, United States</td><td>Snapmaker 2.0</td><td>&amp;nbsp;Model: A250</td></tr><tr><td>3M DP190 Scotch-Weld Gray</td><td>&amp;nbsp;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></tr><tr><td>Ceramic knife</td><td>R felngli, China</td><td>N.A.</td><td>General</td></tr><tr><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 &amp;le; 50 ppm (by K.F.), 99.5%, SafeDry, with molecular sieves, Safeseal</td></tr><tr><td>Hot melt adhesive&amp;nbsp;</td><td>Magic Dragon, China</td><td>N.A.</td><td>JTWJRRJB001</td></tr><tr><td>Inductively Coupled Plasma Mass Spectrometry&amp;nbsp;</td><td>PerkinElmer, Inc., Shelton, CT USA</td><td>N.A.</td><td>Model: NexION 350X</td></tr><tr><td>Medical Infusion Bag&amp;nbsp;</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&amp;Omega;, 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 &amp;lt;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 &amp;lt; 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&amp;nbsp;</td><td>Bond A3 Pipette</td><td>N.A.</td><td>200 &amp;mu;L</td></tr><tr><td>Pipette Tip</td><td>Titan</td><td>T2-H-T0200</td><td>200 &amp;mu;L, 300 &amp;mu;L Tip Box Non-sterile|200 &amp;mu;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

Research

JoVE Journal

Environment

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

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

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

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

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

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

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

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

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

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 ...

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

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

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

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

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

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. ...

The MPLEx Protocol for Multi-omic Analyses of Soil Samples

Mass spectrometry (MS)-based integrated metaproteomic, metabolomic, and lipidomic (multi-omic) studies are transforming our ability to understand and characterize microbial communities in environmental and biological systems. These measurements are even enabling enhanced analyses of complex soil microbial communities, which are the most complex microbial systems known to date. Multi-omic analyses, however, do have sample preparation challenges, since separate extractions are typically needed for each omic study, thereby greatly amplifying the preparation time and amount of sample required. To address this limitation, a 3-in-1 method for the simultaneous extraction of metabolites, proteins, and lipids (MPLEx) from the same soil sample was created by adapting a solvent-based approach. This MPLEx protocol has proven to be both simple and robust for many sample types, even when utilized for limited quantities of complex soil samples. The MPLEx method also greatly enabled the rapid multi-omic measurements needed to gain a better understanding of the members of each microbial community, while evaluating the changes taking place upon biological and environmental perturbations.

A protocol is presented for simultaneously extracting metabolites, proteins, and lipids from a single soil sample, allowing reduced sample preparation times and enabling multi-omic mass spectrometry analyses of samples with limited quantities.

Mass spectrometry (MS)-based integrated metaproteomic, metabolomic, and lipidomic (multi-omic) studies are transforming our ability to understand and ...

Chemistry

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

A Practical and Low-Cost Setup to Measure Methane in Aqueous Samples

Measuring Dissolved Methane in Aquatic Ecosystems Using An Optical Spectroscopy Gas Analyzer

Measuring greenhouse gas (GHG) fluxes and pools in ecosystems are becoming increasingly common in ecological studies due to their relevance to climate change. With it, the need for analytical platforms adaptable to measuring different pools and fluxes within research groups also grows. This study aims to develop a procedure to use portable optical spectroscopy-based gas analyzers, originally designed and marketed for gas flux measurements, to measure GHG concentrations in aqueous samples. The protocol involves the traditional headspace equilibration technique followed by the injection of a headspace gas subsample into a chamber connected through a closed loop to the inlet and outlet ports of the gas analyzer. The chamber is fabricated from a generic mason jar and simple laboratory supplies, and it is an ideal solution for samples that may require pre-injection dilution. Methane concentrations measured with the chamber are tightly correlated (r2 &#62; 0.98) with concentrations determined separately through gas chromatography-flame ionization detection (GC-FID) on subsamples from the same vials. The procedure is particularly relevant for field studies in remote areas where chromatography equipment and supplies are not readily available, offering a practical, cheaper, and more efficient solution for measuring methane and other dissolved greenhouse gas concentrations in aquatic systems.

This study demonstrates an approach to measure methane gas concentrations in aqueous samples using portable optical analyzers coupled to an injection chamber in a closed loop. The results are similar to conventional gas chromatography, presenting a practical and low-cost alternative particularly suitable for remote field studies.

Measuring greenhouse gas (GHG) fluxes and pools in ecosystems are becoming increasingly common in ecological studies due to their relevance to climate ...

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

Summary

Explore More Videos

Chapters in this video

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

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

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

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

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

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

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

The MPLEx Protocol for Multi-omic Analyses of Soil Samples

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

Measuring Dissolved Methane in Aquatic Ecosystems Using An Optical Spectroscopy Gas Analyzer