Separation and Identification of Conventional Microplastics from Farmland Soils

This protocol describe conventional microplastic sampling and sample analyzed from soil. The method includes seven parts. They are soil sampling and preparation, density flotation, impurity digestion, coloration, vacuum filtration, morphological observation, and polymer identification.

Here, we present two different analytical processes of the final two steps, which can be carried out independently of each other depending on instrument availability. Collect a representative soil sample using a five-point sampling method in a double-shaped manner across a steady area. Use a 30-centimeters stainless steel soil auger for collection.

Collect and install the samples in a non-plastic container, for example, aluminum foil. Dry the soil at room temperature away from direct sunlight, or use an oven set to 40 degrees Celsius and dry the soil for a minimum of 24 hours until completely dry. If a soil dryer is available, use it for processing multiple soil samples at the same time as the filter within the individual chambers minimize the risk of cross-contamination.

Once dry, grind the soil if necessary. Use clean, non-plastic implements. Grind and save the dry soil.

Use a two to five-millimeters metal sieve. Using a two-dense small scale, lay out fine plus or minus 0.05 grain of the soil sample onto plastic free weighing paper or aluminum foil. Samples can be stored in plus three containers, for example, glass vials.

Transfer the fine-ground dried soil sample into a clean 600-milliliters glass beaker A.Add 230 milliliters of saturated sodium chloride solution to beaker A.Ensure accurate labeling of all storage containers and beakers. Place the beaker A on a magnetic stirring plate at a glass magnetic stirrer. Stir the solution for 30 minutes at 260 revolutions per minute.

Once fully homogenized, remove the magnetic stirrer from the solution and rinse with saturated sodium chloride solution to prevent plastic particle 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. Once the contents of beaker A have completely separated, carefully transfer the supernatant to a new glass beaker B.Rinse the inner walls of beaker A with saturated sodium chloride solution.

Add four molar sodium hydroxide solution to the sample in beaker B to reach a fixed volume of 500 milliliters. Stir the solution for 30 minutes at 260 revolutions per minute. Then remove the magnetic stirring bar and place the beaker on a flat surface away from direct sunlight and leave it standing overnight.

Once the content of baker B have completely separated. Transfer the supernatant from beaker B to a new beaker C.Rinse inner wall of the beaker b with distilled water to ensure maximum particle transfer. Add the previously prepared new red size solution to beaker C to achieve a final concentration of 0.5 molar.

Stir the solution with glass rod until completely homogenized. Then let the solution incubate for 30 minutes in the dock by covering the beaker with aluminum foil. First, set up the vacuum filtration system as follows.

Glass funnel, metal clamp, vacuum filtration base, collection beaker, connection hose, moisture trap, and vacuum pump. Carefully remove new membranes from its storage container using tweezers. Place the filter membrane centrally and flat on the top of vacuum filtration base.

Ensure secure connection by aligning the vacuum filtration base with a glass funnel, fastening them with a metal clamp. Activate the vacuum filtration and slowly pour the liquid from beaker C into the glass funnel. Rinse beaker C several times with distilled water to maximize particle recovery.

Cover the glass funnel with aluminum foil to minimize contamination. Rinse the side of the glass funnel with distilled water after sample filtration to ensure minimal particle lose. Tear off the vacuum pump and carefully retrieve the filter membrane from the plate using the tweezer.

And place each membranes in an individual glass Petri dish. Add the membranes fully dry before close the Petri dish and wrapping it in aluminum foil. Store it in a dry and dark place until further analyze.

If the exact location of fluorescent particle on the membranes is required for later polymer identification, for example, by using FTIR, please refer to the steps below. Use a black gel pen to gently mark the beginning position 10 marks on the filter membrane following the T shape. Activate the fluorescence instrument as follows, the host, fluorescent sources, monitor, and fluorescence microscope.

Turn on the instrument and set the sources LED up to maximum brightness. Utilize the bright field, the DF, and the fluorescent light, the FL switch button to take DF and FL images respectively. The DP2-BSW software for sample observation recording, but just the microscope definition to make the screen sharper.

Take the bright field pictures under the BF position and turn into FL position and fluorescent filter to take pictures in the dock. Ensure the field of view observation sequence runs from one to 10. Make sure the BF and FL pictures should be taken in the same position.

For polymer identification by using LDIR, perform the microscope steps as below. Set up the microscope system as follows. The camera, filters, magnification and microscope stage, and the computer.

Wrap the filter membrane holders with dust-free tissues. Then secure the membranes in the holder and slide onto the microscope stage. Ensure the camera is connected and the microscope magnification is appropriate for sample type and consistent across all samples from the same set.

To quantify the particles on the recorded images, follow the step-by-step instructions provided in the manuscript. If FTIR is used to identify polymer particles, please refer to the steps below. Turn on the FTIR Spectrometer LUMOS and corresponding software post, for example, observation and recording.

Fill in with liquid nitrogen to active the machine. Clean the probe before mirroring each sample. 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. Measure three to five fixed points on the target fragment, and then position the probe according to the location of this fixed points.

On the result page, save the original data. Solve the spectrum, and compare the spectrum with a plastic spectrum in the standard library to confer the heat quality index of the sample. If LDIR is used for polymer particle identification, follow the steps below.

Place the filter membrane into a new 20-milliliter glass vial. Add 20 milliliters of pure ethanol. Close the vial tightly and wrap the lid with parafilm to prevent leakage.

Sonicate samples in an ultrasonic bath for minimum of one hour until all particles have been resuspended. The membrane may leach color, but this will not interfere with the polymer identification. Remove and discard the membrane.

Place the glass vial with the ethanol solution on a magnetic stirring plant, and add a small magnetic glass stirrer to the vial. Let the ethanol evaporate to less than five milliliters by setting the temperature to 100 degrees Celsius and stirring at a low speed to keep particles suspended. To prepare the sample for analyze on the LDIR, shake the samples slowly until all particles are homogeneously suspended in the solution and quickly prepared 10 microliters of the sample onto the slide and let the ethanol evaporate.

Repeat this step two more times to analyze three replicates per sample on each slide. The LDIR slide is inserted into the instrument and the sample name is entered into the connected software. Subsequently, the instrument initiate an automatic scan.

The resulting analyze provides detailed data on the chemical composition of individual particles, the distribution of different polymers within the sample, as well as particle size. The subsequent data processing is detailed in Section 8 of protocol, for example, by using image J as well as in the calculation section of the result in the manuscript. To validate the recovery range of this methodology, samples from three different solid mattresses, silicone dioxide, the SD, Bentonite Clay, the BT, and soil, were analyzed in sets of three replicates.

Assume that all microplastic particles are uniform sphere. That means per five-gram, dry, solid samples include around more than 48, 740 items. Based on the image J software, the information about the number of particles in a single sample can be reviewed, and this three formulas can be calculated the final recovery rate of the microplastics.

Here are some result of this experiment. First is the recovery rate of microplastic from different solid matrices. The average recovery rates are 84%83%and 90%of BT, SD, and soil respectively.

The interference of result from the blank sample and the chemical identification have been eliminated. On average, 86%of the PE particulates were successfully recovered. The bottom is the result of the polymer type of these samples.

It is shown that except polyethylene, the phenolic resin, polyvinyl chloride, polyamide, and polypropylene are also detected. This result may be contributed to minor sample dose during supernatant transfer, filtration, or incorrect identification. These contaminations could have originated from the filtration devices, laboratory equipment, atmospheric deposition, or distilled water.

There are some pictures were taken with different polymer identification methods. These two pictures are based on the FTIR method, and they are taken in the same area of the membranes on the daylight and the fluorescent light. Particles that appear transparent in figure A while flashing green in figure B are considered likely to be plastic material.

Here is a typical case that shows the comparison of the spectrum between the particle being detected to the standard spectrum diagram. The PE particle spectrum matched against the closest library spectra with some match quality of 98%This picture were taken with the LDIR methods. The real pattern and distribution are showing in figure A and some detailed information like the chemical composition of individual particles.

The match quality ran as well as particle size are shown in Figure B.Microplastic pollution in the terrestrial environment is a scientific topic that have received increasing attention over the last decade. However, only recent microplastic taking soil system have been quantified and the detection method for soil microplastic have not been standardized. This protocol described the methodology for sampling, separation, and the chemical identification of microplastic particle.

To enhance operational ease and the widespread adoption, the method is low-cost and the materials are easily available. This protocol shows potential as a guiding framework, presenting a comprehensive approach suitable for various soil types, ensuring accurate quantification and the analyze of microplastic.