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Method Article

Separation and Identification of Conventional Microplastics from Farmland Soils

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DOI:

10.3791/67064

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March 21st, 2025

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1State Key Laboratory of Nutrient Use and Management, College of Resources and Environmental Sciences; National Academy of Agriculture Green Development, Key Laboratory of Plant-Soil Interactions of Ministry of Education, National Observation and Research Station of Agriculture Green Development (Quzhou, Hebei), China Agricultural University, 2School of Environmental & Natural Sciences, Bangor University, 3College of Resources and Environmental Sciences, Beijing Key Laboratory of Farmland Soil Pollution Prevention and Remediation, China Agricultural University, 4Soil Physics and Land Management Group, Wageningen University & Research, 5Institute of Environment and Sustainable Development in Agriculture, Chinese Academy of Agricultural Sciences, 6College of Land and Environment, Shenyang Agricultural University, 7School of Environmental Science and Engineering, Southern University of Science and Technology, 8Institute of Quality Standard and Testing Technology, Beijing Academy of Agriculture & Forestry Sciences, 9China National Environmental Monitoring Centre, 10Key Laboratory of Prevention and Control of Residual Pollution in Agricultural Film, Ministry of Agriculture and Rural Affairs

In This Article

Summary

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.

Abstract

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 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 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 Fourier-Transform Infrared Spectroscopy (μ-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.

Introduction

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 × 106 to 1.0 × 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 (>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 × 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.

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Protocol

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 µg mL-1) - dissolve 10 mg of Nile Red in 100 mL of appropriate solvent (e.g., methanol, acetone).

1. Soil sampling and preparation

  1. Collect a representative soil sample using a five-point sampling method in a "W" 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.
  2. Dry the soil at room temperature away from direct sunlight.
    1. 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.
    2. Otherwise, use an oven set to 40 ᵒC and dry the soil for a minimum of 24 h, until completely dry.
    3. 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 (> 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.
  3. Using a 2 decimal-place scale, weigh out 5.0 g ± 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).

2. Density flotation

  1. 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.
  2. 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.
  3. 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.

3. Impurity digestion

  1. 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.
  2. 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.
  3. 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.

4. Coloration with Nile Red solution

  1. 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.
    1. 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.
  2. 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.

5. Vacuum filtration

  1. 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 µm 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.
  2. 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.
  3. 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.

6. MPs particle quantification by fluorescence microscopy

  1. 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:
    1. Use a black gel pen to gently mark the beginning position and 10 marks on the filter membrane, following the shape of the "Z" (Figure 2). Place the membrane carefully on the glass slides on the microscope stage using tweezers, ensuring a flat sample surface.
    2. Activate the fluorescence instrument in the following order: the host, fluorescent sources, monitor, and fluorescence microscope. Turn on the instrument and set the source's light knob to maximum brightness. Utilize the bright field (BF) and fluorescent light (FL) switch buttons to take BF and FL images, respectively.
    3. 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.
  2. For polymer identification by using LDIR, perform the microscope steps as below:
    1. 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.
    2. 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.
    3. 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.

7. MPs polymer identification using FTIR or LDIR spectroscopy

  1. If FTIR is used to identify polymer particles, please refer to the steps below.
    1. Turn on the FTIR spectrometer and corresponding software for sample observation and recording. Clean the probe before measuring each sample.
    2. 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.
    3. 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'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 "Z-shaped" 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.
    4. A match is accepted if the hit quality index (HQI) ≥ 0.7. Figure 4 shows an example of particles identified by FTIR spectroscopy in a soil sample spiked with 0.04% PE-MPs.
  2. If LDIR is used for polymer particle identification, follow the steps below:
    1. 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.
    2. 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.
    3. Confirm the successful particle resuspension by reanalyzing membranes under the fluorescent microscope and ensuring particle removal is > 95%.
    4. 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 °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.
    5. 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.
    6. 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 µL 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.
    7. 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.
    8. If the hit quality index (HQI) ≥ 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.

8. Particle quantification of fluorescent membrane images using ImageJ

  1. 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 > Set scale to input image pixel and size (mm) values, then select Global to apply these settings to all images during the session.
  2. Transform the image into a binary image and convert it to 8-bit, using the functions Process > Binary and Image > Type > 8-bit, respectively. To select the parameters for particle analysis, use the functions Analyze > Set measurements and select the desired parameters, e.g., Area, Shape descriptors, and Feret's diameter. Select Add to overlay and determine the number of decimal places for data output.
  3. To analyze particles, select Analyze > Analyze particles and determine the particle size range of interest. Set Circularity to 0-1, unselect Pixel units, enable Overlay 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.

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Results

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 µm). 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 blanks...

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Discussion

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...

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Disclosures

The authors have nothing to disclose.

Acknowledgements

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, "Do agricultural microplastics undermine food security and sustainable development in less economically developed countries?" 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 the International Cooperation and Exchange of the National Natural Science Foundation of China under Grant [NSFC-UNEP: 32261143459].

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
2-decimal balancen/an/aStandard 2-decimal balance
40 °C ovenn/an/aStandard large fan-oven with temperature set to 40 °C
8700 LDIRAgilent Technologiesn/aLDIR used to identify particles
Aluminum containern/an/aStandard aluminum food take-away container
Aluminum foiln/an/aStandard heavy-duty aluminum foil
Axioplan 2Zeissn/aFluorescence microscopy is used to observe microplastic particles in the fluorescent state
Bentonite claySigma Aldrich285234Bentonite clay used for recovery tests
BX53Olympusn/aFluorescence microscopy is used to observe microplastic particles in the fluorescent state
Glass beaker (600 ml)n/an/aStandard glass beaker
Glass bottle (1 l)n/an/aStandard glass bottle
Glass magnetic stirrer barn/an/aStandard glass coated magnetic stirrer bar
Glass measuring cylinder (500 ml)n/an/aStandard glass measuring cylinder
Glass pipette (10 ml)
Glass vacuum filtration devicePyrex (purchased via Sigma Aldrich)SLW5809/KITGlass filtration device with 500 ml funnel, porous plate, and 1 l collection beaker
LUMOS Alpha IIBrukern/aFTIR used to analyze suspect microplastics.
Magnetic stirring platen/an/aStandard magnetic stirring plate
MCE filter membraneJinteng companyJTMF0441/0442White MCE membranes, 0.2 µm pore size, 50 mm diameter, with FTIR method
Nile RedFisher Scientific10464311Nile Red powder used to make stock solution of nile red dye using appropriate solvent (e.g. acetone)
PCTE filter membraneSterlitech Corporation1270060Black PCTE membranes, PVP-free, 0.2 µm pore size, 47 mm diameter, with LDIR method
Silicon dioxideSigma Aldrich18649Silicon dioxide used for recovery tests
Sodium chlorideSigma AldrichS9888Sodium chloride used for density separation
Sodium hydroxideFisher Scientific10675692Sodium hydroxide used for organic matter digestion
Soil augern/an/aLength 30 cm; diameter 2 cm; material stainless steel
Ultra-high molecular weight polyethylene microplastic powderSigma Aldrich434272Polyethylene microplastic used to spike silicon dioxide, bentonite clay, and soil samples for recovery tests
Vacuum pumpVacuubrand GmBH Co KGME 2C NTVacuum pump for vacuum filtration

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Microplastics SeparationSoil MicroplasticsDensity FlotationPolymer IdentificationVacuum FiltrationFluorescent MicroscopySodium Chloride SeparationFourier Transform InfraredLaser Direct InfraredSoil Sampling
Separation and Identification of Conventional Microplastics from Farmland Soils
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