This project was supported by the Science and Technology Major Project of Ordos, China [ZD20232320]; the UKRI Global Challenges Research Fund (GCRF) and the Natural Environment Research Council project, &#34;Do agricultural microplastics undermine food security and sustainable development in less economically developed countries?&#34; under Grant [NE/V005871/1]; and the National Natural Science Foundation of China under Grant [42277097]; the High-level Team Project of China Agricultural University, Professor station of China Agricultural University at Xinzhou Center for Disease Control, and Prevention and Basic Research Program in Xinzhou, Shanxi Province [20230515] and&#160;the International Cooperation and Exchange of the National Natural Science Foundation of China under Grant [NSFC-UNEP: 32261143459].

NOTE: The following solutions need to be prepared at ambient temperature prior to the extraction process: 1) Saturated NaCl solution (5.7 M) - dissolve 1 kg of NaCl in 3 L DI H2O; 2) 4 M NaOH - dissolve 480 g NaOH in 3 L DI H2O; 3) Nile Red (100 &#181;g mL-1) - dissolve 10 mg of Nile Red in 100 mL of appropriate solvent (e.g., methanol, acetone).
1. Soil sampling and preparation
<ol>
	<li>Collect a representative soil sample using a five-point sampling method in a &#34;W&#34; shape manner across the study area (Figure 1). Use a 30 cm stainless-steel soil auger for the collection procedure. Collect and store the samples in a non-plastic container, such as aluminum foil. 
	NOTE: The soil samples can be split into different desired depths (e.g., 0-10, 10-20, and 20-30 cm). To minimize plastic contamination, avoid using plastic sampling equipment and storage containers. Instead, non-plastic containers such as aluminum foil are used to collect and store the samples. These samples can either be bulked to provide one composite sample or kept as independent replicates.</li>
	<li>Dry the soil at room temperature away from direct sunlight.
	<ol>
		<li>If a soil drier is available, use it to process multiple soil samples simultaneously, as the filter within the individual chambers minimizes the risk of cross-contamination.</li>
		<li>Otherwise, use an oven set to 40 &#7506;C and dry the soil for a minimum of 24 h, until completely dry.</li>
		<li>Grind and sieve the dry soil using a 2-5 mm metal sieve. Remove visible plant material, stones, and other inert materials. 
		NOTE: Loosely cover the soil with aluminum foil to minimize airborne contamination within the oven/drier. Collect visible plastic debris (&#62; 5 mm) from the soil with metal tweezers and place them in a plastic-free storage container, if later analysis of macroplastic is to be carried out.</li>
	</ol>
	</li>
	<li>Using a 2 decimal-place scale, weigh out 5.0 g &#177; 0.05 g of the soil sample onto a plastic-free weighing paper or aluminum foil. Use a new weighing paper for different samples to minimize cross-contamination. Samples can be stored in plastic-free containers (e.g., glass vials).</li>
</ol>
2. Density flotation
<ol>
	<li>Transfer the 5.0 g dried soil sample into a clean 600 mL glass beaker (Beaker A). Ensure accurate labeling of all storage containers and beakers.</li>
	<li>Add 230 mL of saturated NaCl solution to beaker A. Place beaker A on a magnetic stirring plate and add a glass magnetic stirrer. Stir the solution for 30 min at 260 rpm, until fully homogenized.</li>
	<li>Once fully homogenized, remove the magnetic stirrer from the solution and rinse with saturated NaCl solution to prevent plastic particles from being carried out of the solution. Place the beaker on a flat surface without direct sunlight and leave it standing overnight, until full density separation has occurred. 
	NOTE: When performing the entire procedure, it is necessary to fully cover with aluminum foil to avoid plastic contaminants from the air.</li>
</ol>
3. Impurity digestion
<ol>
	<li>Once the contents of beaker A have completely separated, carefully transfer the supernatant to a new, clean 600 mL glass beaker (beaker B). Carefully rinse the inner walls of beaker A with saturated NaCl solution. Transfer the supernatant to beaker B. Perform this procedure 2-3 times. 
	NOTE: A total of 200 mL of supernatant is recommended. Ensure that the total supernatant volume is consistent while maintaining the same concentration of digestion solution in the next step.</li>
	<li>Add 4 M NaOH solution to the sample in beaker B to reach a fixed volume of 500 mL. Place beaker B on a magnetic stirring plate, add a glass magnetic stirrer, and stir the solution for 30 min at 260 rpm until fully homogenized. Keep the beaker covered with aluminum foil during this process to minimize airborne contamination.</li>
	<li>Once fully homogenized, remove the magnetic stirrer from the solution and rinse with saturated NaCl solution to remove any attached particles. Place beaker B on a flat surface without direct sunlight and leave it standing overnight until full density separation and organic matter digestion have occurred. 
	NOTE: When performing the entire procedure, it is necessary to fully cover with aluminum foil to avoid plastic contaminants entering the solution from the air. The digestion duration depends on the quantity and the type of organic material. Extend the digestion time if required to ensure complete organic matter digestion. After successful digestion, the supernatant should look clear with no visible organic matter floating in the beaker.</li>
</ol>
4. Coloration with Nile Red solution
<ol>
	<li>Once the contents of beaker B have completely separated, carefully transfer the supernatant to a new, clean 600 mL glass beaker (beaker C). Rinse the inner walls of beaker B with DI water to ensure maximum particle transfer.
	<ol>
		<li>If the volume in beaker C is less than 500 mL, make up the value to 500 mL with DI water to unify the solution volume.</li>
	</ol>
	</li>
	<li>Add the Nile Red solution to beaker C to achieve a final maximum concentration of 0.5 M. Stir the solution with a glass rod until completely homogenized, then let the solution incubate for 30 min in the dark by covering the beaker with aluminum foil.</li>
</ol>
5. Vacuum filtration
<ol>
	<li>Set up the vacuum filtration devices in the following order: glass funnel, metal clamp, vacuum filtration base, collection beaker, connection hose, moisture trap, and vacuum pump. Carefully remove a new membrane (0.2 &#181;m pore size, 47 mm diameter) from its storage container using metal tweezers. Place the filter membrane centrally and flat on the top of the vacuum filtration base. 
	NOTE: Ensure a secure connection by aligning the vacuum filtration base with the glass funnel and fastening it with a metal clamp.</li>
	<li>Activate the vacuum filtration and slowly pour the liquid from beaker C into the glass funnel. Rinse beaker C several times with DI water to maximize particle recovery. Rinse the sides of the glass funnel with DI water after sample filtration to ensure minimal particle loss. 
	NOTE: Cover the glass funnel with aluminum foil to minimize airborne contamination during the filtration process. If the sample has a high particle number and the filtration speed decreases, multiple membranes can be used for the same sample. This will ensure an even distribution of particles across the membrane and minimize the risk of particles aggregating and overlapping for later quantification.</li>
	<li>Once filtration is completed, carefully retrieve the filter membrane from the porous plate using tweezers and place each membrane in an individual glass Petri dish. Let the membrane fully dry before closing the Petri dish and wrapping it in aluminum foil. Store it in a dry and dark place until further analysis.</li>
</ol>
6. MPs particle quantification by fluorescence microscopy
<ol>
	<li>If the exact location of fluorescent particles on the membrane is required for later polymer identification (e.g., by using FTIR), please refer to the steps below:
	<ol>
		<li>Use a black gel pen to gently mark the beginning position and 10 marks on the filter membrane, following the shape of the &#34;Z&#34; (Figure 2). Place the membrane carefully on the glass slides on the microscope stage using tweezers, ensuring a flat sample surface.</li>
		<li>Activate the fluorescence instrument in the following order: the host, fluorescent sources, monitor, and fluorescence microscope. Turn on the instrument and set the source&#39;s light knob to maximum brightness. Utilize the bright field (BF) and fluorescent light (FL) switch buttons to take BF and FL images, respectively.</li>
		<li>Using the software for sample observation and recording (e.g., DP2-BSW), take bright field pictures under the BF position. Turn the knob into FL position and take pictures with the fluorescence filter in the dark. Ensure the field of view observation sequence runs from 1 to 10. Make sure the FL and BL pictures are taken in the same position. 
		NOTE: Fluorescent microscopic analysis should be carried out within 24-48 h after filtration to ensure optimal fluorescence of particles. Ensure the BF and FL images are taken in the same position (Figure 3). Ensure all membranes are analyzed using the same magnification and instrument settings.</li>
	</ol>
	</li>
	<li>For polymer identification by using LDIR, perform the microscope steps as below:
	<ol>
		<li>Set up the microscopy system as follows: the camera, filters, magnifications and microscope stage, and computer. Select the appropriate filter mode best suited for the desired excitation and emission wavelengths (e.g., 470 nm and 495 nm, respectively) depending on sample and background fluorescence.</li>
		<li>Wipe the filter membrane holders with dust-free tissues, then secure the membrane in the holder and slide onto the microscope stage. Scan the entire membrane visually to ensure an even distribution of particles.</li>
		<li>Ensure the camera is connected, and the microscope magnification is appropriate for all sample types and consistent across all samples from the same set. Determine the size of the area in mm captured with the camera before starting the analysis by using a measuring tape. 
		NOTE: If particles are evenly distributed, analyze a minimum of 10% of the total membrane area by selecting the required number of sampling points on the membrane. Take a picture of each sampling point using the camera.</li>
	</ol>
	</li>
</ol>
7. MPs polymer identification using FTIR or LDIR spectroscopy
<ol>
	<li>If FTIR is used to identify polymer particles, please refer to the steps below.
	<ol>
		<li>Turn on the FTIR spectrometer and corresponding software for sample observation and recording. Clean the probe before measuring each sample.</li>
		<li>Identify the particles for monitoring through real-time screen recording. Adjust position and sharpness by manipulating the rocker. Bring the operating platform to the center and capture the current air background spectrum. 
		NOTE: Target particles that correspond to both BF and FL images.</li>
		<li>Measure 3-5 fixed points on each particle to attain spectra within the infrared wavenumber range of 400-4,000 cm-1. On the results page, save the original data, obtain the spectrum, and compare it with the plastic spectrum in the standard library to confirm the sample&#39;s hit quality index (Figure 4). 
		NOTE: In this study, due to the small size of MPs and the challenges in quantifying them across the entire filter, 5-7 representative particles were selected from each sample for FTIR analysis. To ensure an even distribution, particles following a &#34;Z-shaped&#34; pattern across the filter were selected. However, it should be noted that some uncertainty may arise, as the particles were selected randomly rather than by scanning the entire filter.</li>
		<li>A match is accepted if the hit quality index (HQI) &#8805; 0.7. Figure 4 shows an example of particles identified by FTIR spectroscopy in a soil sample spiked with 0.04% PE-MPs.</li>
	</ol>
	</li>
	<li>If LDIR is used for polymer particle identification, follow the steps below:
	<ol>
		<li>Once the fluorescent microscope steps are completed, resuspend the particles for analysis on the LDIR. Place the filter membrane into a new glass vial and add 20 mL of pure ethanol. Close the vials tightly and wrap the lids with paraffin film to prevent leakage.</li>
		<li>Sonicate samples in an ultrasonic bath for a minimum of 1 h until all particles have been resuspended. Remove and discard the membrane from the glass vial. 
		NOTE: Ensure the membrane side with particles faces inwards, i.e., away from the vial wall. Sonication time depends on the membrane type used; the membrane may leach color, but this will not interfere with the polymer identification.</li>
		<li>Confirm the successful particle resuspension by reanalyzing membranes under the fluorescent microscope and ensuring particle removal is &#62; 95%.</li>
		<li>Place the glass vials with the ethanol solution on a magnetic stirring plate and add a small magnetic glass stirrer to the vial. Let the ethanol evaporate to less than 5 mL by setting the plate temperature to 100 &#176;C and stirring at a low speed to keep particles suspended. 
		NOTE: Cover the vials lightly with aluminum foil to minimize airborne contamination of the samples. Optionally dry the solution under a gentle stream of nitrogen for ethanol evaporation.</li>
		<li>Transfer the sample into a small 8 mL glass vial, using new ethanol to rinse the 20 mL glass vial to maximize particle recovery. 
		NOTE: If required, add new ethanol to the small glass vial to reach exactly 5 mL.</li>
		<li>To prepare the sample for analysis on the LDIR, shake the sample thoroughly until all particles are homogenously suspended in the solution and quickly pipette 10 &#181;L of the sample onto the slide and let the ethanol evaporate. Repeat this step two more times to analyze 3 replicates per sample on each slide.</li>
		<li>Use the particle analysis tool in the associated software to measure the infrared spectra of each particle within the set size range and match them against the internal library for polymer identification. 
		NOTE: One point on each particle is automatically measured in the infrared wavenumber range of 800-1800 cm-1, and particles are automatically matched against known library spectra of polymers and other organic and inorganic materials.</li>
		<li>If the hit quality index (HQI) &#8805; 0.8, the match is accepted. Figure 5 shows an example of particles identified by LDIR spectroscopy in a soil sample spiked with 0.04% PE-MPs. 
		NOTE: Sample volume can be increased for samples with low particle numbers.</li>
	</ol>
	</li>
</ol>
8. Particle quantification of fluorescent membrane images using ImageJ
<ol>
	<li>Open ImageJ32 (version 1.54f) and load images into the software. Adjust the scale of the image to the correct measurements corresponding to the real image size. Use the functions Analyze &#62; Set scale to input image pixel and size (mm) values, then select Global to apply these settings to all images during the session.</li>
	<li>Transform the image into a binary image and convert it to 8-bit, using the functions Process &#62; Binary and Image &#62; Type &#62; 8-bit, respectively. To select the parameters for particle analysis, use the functions Analyze &#62; Set measurements and select the desired parameters, e.g., Area, Shape descriptors, and Feret&#39;s diameter. Select Add to overlay and determine the number of decimal places for data output.</li>
	<li>To analyze particles, select Analyze &#62; Analyze particles and determine the particle size range of interest. Set Circularity to 0-1, unselect Pixel units, enable Overlay&#160;to be shown, and select the following boxes: Display results, Exclude edges, Clear results, Include holes, Summarize, and Overlay. Export results as .csv files from the results window. 
	NOTE: The analysis of fluorescent images will only provide information on particle number, not polymer number; therefore, an additional identification procedure is required, for example, using IR spectroscopy techniques.</li>
</ol>

The authors have nothing to disclose.

The soil sampling strategy in the field, including approaches such as simple random sampling or systematic grid sampling, as well as the sampling area and depth, must be tailored to the specific research questions and clearly defined prior to the sample collection. Some studies have focused on the topsoil layer of 0-10 cm34,37, whereas other collected soil samples with a depth of 0-40 cm38. Since the size and abundance of MPs vary in diffe...

It is estimated that 4.8 to 12.7 million metric tons of plastic enter the ocean annually from terrestrial sources1,2. These plastic particles gradually degrade into smaller fragments in response to ultraviolet irradiation, mechanical abrasion, and biodegradation3,4. Microplastics (MPs) pollution, with plastic particles of diameter less than 5 mm, in the soil is becoming an increasing concern, particularly in terms of its potential effect on soil and crop health. It is primarily driven by the continual rise in plastic production and challenges surrounding the appropriate disposal of plastic waste5,6.
The accumulation of MPs in soil can be attributed to various external factors. The potential sources of MPs in soils are complex, including the utilization of plasticulture practices (e.g., plastic mulch films, irrigation pipes, greenhouse films, and associated infrastructure)7,8,9 and input of organic amendments (such as sewage sludge application, agricultural compost, and organic fertilizer)10. In addition, the inappropriate disposal of plastic litter11, decomposition of digested food waste from food plastic packaging residue12, utilization of coated fertilizers13, wear and tear of rubber tires14, and atmospheric deposition15 are also known contributors to MPs in soils. China, the leading producer and user of agricultural plastics, particularly plastic mulch films, has been estimated to have an average abundance of MPs in heavily plastic-mulched agricultural farmland of ca. 4231 items kg-1 (dry soil)16. In 2018, the quantities of MPs in Chinese farmland soils within the 0-10 cm depth ranged from 4.9 &#215; 106 to 1.0 &#215; 107 tons, with a significant contribution from agricultural mulch films17. Sludge applications to agricultural soils in Europe and North America may input over 63,000 and 44,000 tons of MPs per year, respectively18. A study in Germany showed that compost applications to arable fields also led to an annual input of plastic particles (&#62;1 mm) into arable fields. The application of compost led to 35 billion to 2.2 trillion plastic particles10.The contribution of atmospheric MPs to soils is still uncertain and requires further quantification15. For example, the annual average input of atmospheric MPs is estimated to be 7.9 &#215; 104 items m-2 yr-1 in China16. The extremely wide range of sources of MPs in soil has attracted the attention of many researchers, but due to the diversity of sampling, extraction, and analytical detection methods, it is difficult to integrate and compare the results of various studies.
The accumulation of MPs from a wide range of sources poses a potential environmental threat to global soils16, highlighting the clear need for studies of MPs in soil. Some studies have shown the effects of MPs on agricultural soil include altering soil properties, impeding the growth and development of plants and soil organisms, and impacting soil microbial activity19,20. Other studies have found that MPs can accumulate in organisms at higher trophic levels along the food chain21, leading to a potential hazard to human health22. To clarify the soil environmental effects of MPs, it is first necessary to understand the current status of their contamination, including their abundance, polymer identification, and distribution characteristics. Therefore, the accurate identification and detection of soil MPs are of paramount importance.
Currently, a growing number of articles are exploring the global presence of MPs in soil, with considerable variation observed in the extraction and detection methods23. After the careful collection of samples (to minimize MPs contamination), the protocol for MPs analysis typically involves three key steps. First, density separation is widely adopted to isolate MPs particles from the soil matrix. This process commonly utilizes reagents such as distilled (DI) water (1.0 g cm-3), sodium chloride (NaCl, 1.2 g cm-3), or zinc chloride (ZnCl2, 1.6 g cm-3). Secondly, methods for removing organic impurities from the surface of MPs include cleaning with acidic and alkaline solutions or other oxidizing agents and enzymatic digestion24. The digestion of organic matter in the soil matrix or adhering to MPs particles is commonly carried out using 30% hydrogen peroxide (H2O2), 65% nitric acid (HNO3), or 50% sodium hydroxide (NaOH)25. Following the density separation and organic matter digestion, the microscopic examination of MPs samples is required to determine the number of particles. This examination is supplemented with the analysis of the chemical composition of the polymers through techniques such as Fourier-Transform Infrared Spectroscopy (FTIR), Raman spectroscopy, or other near-infrared spectroscopy techniques26.
However, each step in the MPs extraction and detection process carries the potential for either overestimation or underestimation of MPs occurrence. For instance, despite the widespread use of DI water as a reagent for density separation due to its cost-effectiveness and lack of hazardous properties, it may lead to the exclusion of MPs particles with higher density27. Conversely, the widespread application of high-density reagents may be limited by environmental hazards and increased costs28. Additionally, certain reagents used for organic digestion have the potential to cause damage to MPs particles29. Furthermore, visual classification using optical, stereoscopic, and anatomical microscopy is not without its challenges26,30. The determination of MPs particles heavily relies on the expertise and operation of the analysts, as well as the instrument settings. These findings emphasize the difficulty in achieving consistency and accuracy when employing various methodologies, thereby complicating the comparison of results across different studies.
To ensure the reliability and comparability of data across studies, it is imperative to establish a standardized protocol for MPs extraction and detection in soil. This standardization will not only enhance the accuracy of MPs occurrence assessments but also facilitate a more comprehensive and unified understanding of the environmental impact of MPs in soil ecosystems. To address the limitations of extraction and detection methods, the selected reagents for standardized methods should be readily available, should not affect the integrity or chemical composition of the MPs particles, and pose the lowest feasible environmental risk. Moreover, standardized methods should demonstrate high efficiency in both recovering MPs and removing organic matter from the soil matrix.
An easy-to-follow protocol is vital for widespread adoption across different research settings. Considering both MPs recovery rates and cost-effectiveness, saturated NaCl is the optimal choice for large-scale soil sample density separation. For the digestion of organic matter, NaOH was used, as preliminary isolation experiments have shown that 4 M NaOH solution effectively decomposes soil sample impurities, such as plant residues, without causing significant damage to the MPs. In general, this experimental method utilizes readily available and cost-effective materials, has low operational complexity, and ensures a reliable extraction rate.
We recommend using the rapid and economical separation methodology proposed by Chinese Academy of Agricultural Sciences to determine MPs collected in agricultural fields31. For all following steps, ensure that all containers, instruments, and glassware are cleaned with DI water before use to minimize contamination. Also, ensure to run regular blanks alongside the samples to account for contamination introduced by the collection and extraction procedures.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>2-decimal balance</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard 2-decimal balance</td>
		</tr>
		<tr>
			<td>40 &#176;C oven</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard large fan-oven with temperature set to 40 &#176;C</td>
		</tr>
		<tr>
			<td>8700 LDIR</td>
			<td>Agilent Technologies</td>
			<td>n/a</td>
			<td>LDIR used to identify particles</td>
		</tr>
		<tr>
			<td>Aluminum container</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard aluminum food take-away container</td>
		</tr>
		<tr>
			<td>Aluminum foil</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard heavy-duty aluminum foil</td>
		</tr>
		<tr>
			<td>Axioplan 2</td>
			<td>Zeiss</td>
			<td>n/a</td>
			<td>Fluorescence microscopy is used to observe microplastic particles in the fluorescent state</td>
		</tr>
		<tr>
			<td>Bentonite clay</td>
			<td>Sigma Aldrich</td>
			<td>285234</td>
			<td>Bentonite clay used for recovery tests</td>
		</tr>
		<tr>
			<td>BX53</td>
			<td>Olympus</td>
			<td>n/a</td>
			<td>Fluorescence microscopy is used to observe microplastic particles in the fluorescent state</td>
		</tr>
		<tr>
			<td>Glass beaker (600 ml)</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard glass beaker</td>
		</tr>
		<tr>
			<td>Glass bottle (1 l)</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard glass bottle</td>
		</tr>
		<tr>
			<td>Glass magnetic stirrer bar</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard glass coated magnetic stirrer bar</td>
		</tr>
		<tr>
			<td>Glass measuring cylinder (500 ml)</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard glass measuring cylinder</td>
		</tr>
		<tr>
			<td>Glass pipette (10 ml)</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glass vacuum filtration device</td>
			<td>Pyrex (purchased via Sigma Aldrich)</td>
			<td>SLW5809/KIT</td>
			<td>Glass filtration device with 500 ml funnel, porous plate, and 1 l collection beaker</td>
		</tr>
		<tr>
			<td>LUMOS Alpha II</td>
			<td>Bruker</td>
			<td>n/a</td>
			<td>FTIR used to analyze suspect microplastics.</td>
		</tr>
		<tr>
			<td>Magnetic stirring plate</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Standard magnetic stirring plate</td>
		</tr>
		<tr>
			<td>MCE filter membrane</td>
			<td>Jinteng company</td>
			<td>JTMF0441/0442</td>
			<td>White MCE membranes, 0.2 &#181;m pore size, 50 mm diameter, with FTIR method</td>
		</tr>
		<tr>
			<td>Nile Red</td>
			<td>Fisher Scientific</td>
			<td>10464311</td>
			<td>Nile Red powder used to make stock solution of nile red dye using appropriate solvent (e.g. acetone)</td>
		</tr>
		<tr>
			<td>PCTE filter membrane</td>
			<td>Sterlitech Corporation</td>
			<td>1270060</td>
			<td>Black PCTE membranes, PVP-free, 0.2 &#181;m pore size, 47 mm diameter, with LDIR method</td>
		</tr>
		<tr>
			<td>Silicon dioxide</td>
			<td>Sigma Aldrich</td>
			<td>18649</td>
			<td>Silicon dioxide used for recovery tests</td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sigma Aldrich</td>
			<td>S9888</td>
			<td>Sodium chloride used for density separation</td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Fisher Scientific</td>
			<td>10675692</td>
			<td>Sodium hydroxide used for organic matter digestion</td>
		</tr>
		<tr>
			<td>Soil auger</td>
			<td>n/a</td>
			<td>n/a</td>
			<td>Length 30 cm; diameter 2 cm; material stainless steel</td>
		</tr>
		<tr>
			<td>Ultra-high molecular weight polyethylene microplastic powder</td>
			<td>Sigma Aldrich</td>
			<td>434272</td>
			<td>Polyethylene microplastic used to spike silicon dioxide, bentonite clay, and soil samples for recovery tests</td>
		</tr>
		<tr>
			<td>Vacuum pump</td>
			<td>Vacuubrand GmBH Co KG</td>
			<td>ME 2C NT</td>
			<td>Vacuum pump for vacuum filtration</td>
		</tr>
	</tbody>
</table>

separation and identification of conventional microplastics from farmland soils

Microplastics (MPs) pollution in the terrestrial environment has received increasing attention over the last decade, with increasing studies describing the numbers and types of MPs in different soil systems and their impacts on soil and crop health. However, different MPs extraction and analytical methods are used, limiting opportunities to compare results and generate reliable evidence for industry advice and policymakers. Here, we present a protocol that describes the methodology for sampling, separation, and chemical identification of conventional MPs&#160;from soil. The method is low-cost, and the materials are readily available. This enhances operational ease and may help with widespread adoption. The protocol provides detailed information on sample collection from the top 0-30 cm of soil using plastic-free utensils; simulation of different soil types through the use of various solid media (such as bentonite clay, silicon dioxide, and non-contaminated soil), with the addition of the same mass&#160;of polyethylene(PE)-MPs for subsequent quantification; density separation of plastic particles utilizing saturated sodium chloride (NaCl) solution and digestion of organic impurities in the supernatant using 4 M sodium hydroxide (NaOH) solution; quantification of particles using fluorescent microscopy after Nile Red staining; and polymer identification using micro&#160;Fourier-Transform Infrared Spectroscopy (&#956;-FTIR) or Laser-Direct Infrared (LDIR) spectroscopy. The MPs recovery rate ranged from 83% - 90% for the abovementioned media. This protocol presents an efficient method for soil MPs analysis that is optimized for feasibility, applicability, and cost-effectiveness. Moreover, the video accompanied can guide the process of analyzing the soil MPs step-by-step virtually. This study is dedicated to standardizing the methods for soil MPs analysis, enhancing the connectivity and comparability of measurements, and establishing a foundation for more standardized and scientific research.

To validate the recovery rates of this methodology, samples from three different solid matrices (silicon dioxide (SD), bentonite clay (BT), and soil) were analyzed in sets of three replicates. Samples were analyzed with and without the addition of 0.04% w/w white polyethylene (PE) microplastic (particle size range 40-48 &#181;m). Soil samples were collected from Haidian District, Beijing, China (China Agricultural University West Campus), and soil was classified as umber soil. Additionally, three replicates of b...

Presented here is a method for extracting microplastics from soil and identifying their polymer types. The method has been optimized for execution, applicability, and cost-effectiveness. It lays a scientific foundation for standardizing the analytical method to identify microplastics in soils.

Separation and Identification of Conventional Microplastics from Farmland Soils

Microplastics (MPs) pollution in the terrestrial environment has received increasing attention over the last decade, with increasing studies describing ...

separation-identification-conventional-microplastics-from-farmland

Research

JoVE Journal

Environment

852 Views.  China Agricultural University. Presented here is a method for extracting microplastics from soil and identifying their polymer types. The method has been optimized for execution, applicability, and cost-effectiveness. It lays a scientific foundation for standardizing the analytical method to identify microplastics in soils.

Video: Separation and Identification of Conventional Microplastics from Farmland Soils

Helminth Collection and Identification from Wildlife

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), &#34;thorny-headed worms&#34; (acanthocephalans), &#34;flukes&#34; (trematodes), and &#34;tapeworms&#34; (cestodes). The optimum method for collecting helminths is to examine a host that has been dead less than 4-6 hr since most helminths will still be alive. A thorough necropsy should be conducted and all major organs examined. Organs are washed over a 106 &#956;m sieve under running water and contents examined under a stereo microscope. All helminths are counted and a representative number are fixed (either in 70% ethanol, 10% buffered formalin, or alcohol-formalin-acetic acid). For species identification, helminths are either cleared in lactophenol (nematodes and small acanthocephalans) or stained (trematodes, cestodes, and large acanthocephalans) using Harris&#39; hematoxylin or Semichon&#39;s carmine. Helminths are keyed to species by examining different structures (e.g. male spicules in nematodes or the rostellum in cestodes). The protocols outlined here can be applied to any vertebrate animal. They require some expertise on recognizing the different organs and being able to differentiate helminths from other tissue debris or gut contents. Collection, preservation, and staining are straightforward techniques that require minimal equipment and reagents. Taxonomic identification, especially to species, can be very time consuming and might require the submission of specimens to an expert or DNA analysis.

Wild animals are commonly parasitized by a wide range of helminths.&#160; The four major types of helminths are &#8220;roundworms&#8221; (nematodes), &#8220;thorny-headed worms&#8221; (acanthocephalans), &#8220;flukes&#8221; (trematodes), and &#8220;tapeworms&#8221; (cestodes).&#160; Here we describe how helminths are collected from a vertebrate animal and how they are preserved and taxonomically identified.&#160;

Wild animals are commonly parasitized by a wide range of helminths. The four major types of helminths are &#34;roundworms&#34; (nematodes), ...

Measurement of Greenhouse Gas Flux from Agricultural Soils Using Static Chambers

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

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

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

A Bending Test for Determining the Atterberg Plastic Limit in Soils

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

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

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

Extraction and Analysis of Microbial Phospholipid Fatty Acids in Soils

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

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

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

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

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

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

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

Protocol for Microplastics Sampling on the Sea Surface and Sample Analysis

Microplastic pollution in the marine environment is a scientific topic that has received increasing attention over the last decade. The majority of scientific publications address microplastic pollution of the sea surface. The protocol below describes the methodology for sampling, sample preparation, separation and chemical identification of microplastic particles. A manta net fixed on an &#187;A frame&#171; attached to the side of the vessel was used for sampling. Microplastic particles caught in the cod end of the net were separated from samples by visual identification and use of stereomicroscopes. Particles were analyzed for their size using an image analysis program and for their chemical structure using ATR-FTIR and micro FTIR spectroscopy. The described protocol is in line with recommendations for microplastics monitoring published by the Marine Strategy Framework Directive (MSFD) Technical Subgroup on Marine Litter. This written protocol with video guide will support the work of researchers that deal with microplastics monitoring all over the world.

The protocol below describes the methodology for: microplastics sampling on the sea surface, separation of microplastic and chemical identification of particles. This protocol is in line with the recommendations for microplastics monitoring published by the MSFD Technical Subgroup on Marine Litter.

Microplastic pollution in the marine environment is a scientific topic that has received increasing attention over the last decade. The majority of ...

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 ...

Combined Size and Density Fractionation of Soils for Investigations of Organo-Mineral Interactions

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF relies on sequential density separation and sedimentation steps to isolate (1) the free light fraction (uncomplexed organic matter), (2) the occluded light fraction (uncomplexed organic matter trapped in soil aggregates) and (3) a variable number of heavy fractions (soil minerals and their associated organic matter) differing in composition. Provided that the parameters of the CSDF (dispersion energy, density cut-offs, sedimentation time) are properly selected, the method yields heavy fractions of relatively homogeneous mineral composition. Each of these fractions is expected to have a different complexing ability towards organic matter, rendering this a useful method to isolate and study the nature of organo-mineral interactions. Combining density and particle size separation brings an improved resolution compared to simple size or density fractionation methods, allowing the separation of heavy components according to both mineralogy and size (related to surface area) criteria. As is the case for all physical fractionation methods, it may be considered as less disruptive or aggressive than chemically-based extraction methods. However, CSDF is a time-consuming method and furthermore, the quantity of material obtained in some fractions can be limiting for subsequent analysis. Following CSDF, the fractions may be analyzed for mineralogical composition, soil organic carbon concentration and organic matter chemistry. The method provides quantitative information about organic carbon distribution within a soil sample and brings light to the sorptive capacity of the different, naturally-occurring mineral phases, thus providing mechanistic information about the preferential nature of organo-mineral interactions in soils (i.e., which minerals, what type of organic matter).

Combined size and density fractionation (CSDF) is a method to physically separate soil into fractions differing in texture (particle size) and mineralogy (density). The purpose is to isolate fractions with different reactivities towards soil organic matter (SOM), in order to better understand organo-mineral interactions and SOM dynamics.

Combined size and density fractionation (CSDF) is a method used to physically separate soil into fractions differing in particle size and mineralogy. CSDF ...

Chemical Isolation, Quantification, and Separation of Skin Lipids from Reptiles

Reptiles signal to conspecifics using lipids in their skin, primarily to enable mate tracking and assessment. The isolation of these lipids has utility in research focused on evolutionary patterns and mechanisms of chemical communication, in addition to understanding the waterproofing role of lipids in the evolution of terrestrial life. In an applied approach, such skin-based cues have potential use for wildlife managers dealing with invasive species. The main steps for quantifying reptile skin lipids in the protocol presented here include extraction, total lipid determination, and fractionation via column chromatography, the latter process resulting in purified eluates of compounds which can then either be analyzed to assign compound identifications (e.g., gas chromatography-mass spectrometry [GC-MS]) and/or used directly in more refined bioassays. Skin lipids can be extracted from living skin, shed skin, or dead whole animals, using nonpolar organic solvents (e.g., hexane, benzene, toluene). Extraction solubilizes the lipids and, then, the solvent can be evaporated to yield a measurable lipid-only extract. Fractionation involves the separation of the total lipid extract into specific eluates via traditional column chromatography. The total lipid extract is first bound to a substrate-based column (e.g., alumina) and, then, individual eluates (&#34;fractions&#34;) of solvent at specific volumes are passed sequentially through the column to elute sets of compounds from the lipid mixture based on common polarity. The fractions progress in polarity at a standardized sequence by increasing the relative amount of polar solvent (e.g., diethyl ether) in nonpolar solvent. In this manuscript, we describe several methods for extracting skin lipids of reptiles and, then, provide a standard protocol for isolating different sets of compounds based on polarity, using traditional column chromatography. Whole lipid extracts or specific fractions can, then, be used in bioassays to determine any biological activity elicited by the compounds therein.

In reptiles, skin lipids from conspecifics are crucial for sexual signaling, with potential use in invasive species management. Here, we describe protocols for extracting skin lipids from shed skin or whole animals, determining and analyzing the total lipid mass, and separating the lipids using fractionation via column chromatography.

Reptiles signal to conspecifics using lipids in their skin, primarily to enable mate tracking and assessment. The isolation of these lipids has utility in ...

High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling in soils, to pathogen competition, plant growth promotion, and cross-kingdom signaling. Furthermore, siderophores are also of commercial interest in bioleaching and bioweathering of metal-bearing minerals and ores. A rapid, cost effective, and robust means of quantitatively assessing siderophore production in complex samples is key to identifying important aspects of the ecological ramifications of siderophore activity, including, novel siderophore producing microbes. The method presented here was developed to assess siderophore activity of in-tact microbiome communities, in environmental samples, such as soil or plant tissues. The samples were homogenized and diluted in a modified M9 medium (without Fe), and enrichment cultures were incubated for 3 days. Siderophore production was assessed in samples at 24, 48, and 72 hours (h) using a novel 96-well microplate CAS (Chrome azurol sulphonate)-Fe agar assay, an adaptation of the traditionally tedious and time-consuming colorimetric method of assessing siderophore activity, performed on individual cultivated microbial isolates. We applied our method to 4 different genotypes/Lines of wheat (Triticum aestivum L.), including Lewjain, Madsen, and PI561725, and PI561727 commonly grown in the inland Pacific Northwest. Siderophore production was clearly impacted by the genotype of wheat, and in the specific types of plant tissues observed. We successfully used our method to rapidly screen for the influence of plant genotype on siderophore production, a key function in terrestrial and aquatic ecosystems. We produced many technical replicates, yielding very reliable statistical differences in soils and within plant tissues. Importantly, the results show the proposed method can be used to rapidly examine siderophore production in complex samples with a high degree of reliability, in a manner that allows communities to be preserved for later work to identify taxa and functional genes.

We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling ...