The project for which this protocol was established was funded by the National Oceanic and Atmospheric Administration (NOAA) Marine Debris Program (# NA16NO29990029). We thank Miles Corcoran at the National Great Rivers Research and Education Center (NGRREC) in Alton, Illinois, for help with site selection and boat operation. Field and lab work was completed with the help of Camille Buckley, Michael Abegg, Josiah Wray, and Rebecca Wagner.

1. Water Sample Collection
<ol>
	<li>Collect water samples and water quality data of interest by boat where the river is well-mixed, ideally at locations where river stage or discharge is known (e.g., United States Geological Survey (USGS) gauging stations).20 To assure that the water is well-mixed, guide the boat using a handheld meter immersed in the river to where conductivity stays relatively constant.</li>
	<li>At the sampling sites, record location coordinates and depth. To find the 0.6-depth, simply multiply the total depth by 0.6. Measure water quality parameters of interest (e.g., turbidity, temperature, conductivity, pH, and dissolved oxygen (DO)) using a handheld meter. To measure the parameters, pump sample water from the desired depth into a wide-mouth container using the peristaltic pump and immediately take the measurements (step 1.5).</li>
	<li>Use a peristaltic pump with tubing to obtain samples from the surface and 0.6-depth. Attach the correct tubing length to the pump for the given depth.
	<ol>
		<li>Due to the strong currents in river systems, attach a 6.4 mm welded chain to the pump tubing using zip ties to help weight the tubing. At the end of the chain, place a weight or cement block to further weight the chain and tubing assembly. 
		Caution: Do not attach the weight or cement block directly to the pump tubing.</li>
	</ol>
	</li>
	<li>Place the effluent end of the tubing over the boat&#39;s edge, away from clothing that could shed fibers. Slowly lower the influent end of the tubing to the desired depth (i.e., the surface or 0.6-depth). Then, run the pump in reverse to purge the tubing with air for at least 30 s. After air purging, reverse the pump direction and rinse the tubing with sample water from the desired depth while allowing the water to drain off the boat or into a waste container. Stop the pump after the tubing has been rinsed for at least 30 s.</li>
	<li>Rinse the container used for water quality measurements three times with sample water, dumping the rinse water each time. Once rinsed, fill the container with sample water and measure the water quality parameters of interest using a handheld meter (step 1.2).</li>
	<li>Collect a microplastic subsample by placing the tubing effluent into a labeled, 1 L container that has been pre-rinsed with at least 250 mL of DI water three times. Then, rinse the container three additional times with the sample water, discarding the rinse water each time. Once the microplastic container is rinsed, fill it with the sample.</li>
	<li>Using the same peristaltic pump method outlined in step 1.6, collect a subsample for total suspended solids (TSS) in a labeled, 250 mL bottle that has been pre-rinsed with at least 100 mL of DI water three times. Rinse the bottle three more times with sample water, discarding the rinse water each time. Once the TSS container is rinsed, fill it with the sample.</li>
	<li>Collect field triplicates and blanks at least once per day in the field, in the same manner described in steps 1.6-1.7, for quality assurance/quality control (QA/QC) purposes. To collect a blank, bring two 1 L containers of DI water to the field. After purging the pump tubing with air, open the first container of DI water and rinse the pump tubing using the method described in step 1.4. Once the tubing is rinsed, open the second container of DI water and pump it into an empty 1 L container and a 250 mL bottle for microplastic and TSS blanks, respectively.</li>
	<li>Store the microplastic and TSS subsamples on ice until returning to the lab, where they will be stored at -20&#176;C until they are processed. 
	Caution: Make sure to leave some head space in the sample containers so that they are not damaged due to ice expansion when freezing. 
	Note: The protocol can be paused here.</li>
</ol>
2. TSS Determination
<ol>
	<li>Use United States Environmental Protection Agency (USEPA) method 160.2 to determine TSS with the 250 mL subsamples collected in the field23. Compare the calculated TSS values with the total plastics found.</li>
</ol>
3. Microplastic Filtration Device Assembly
<ol>
	<li>Thoroughly rinse the filtration device and nylon mesh sieves (Figure 1) three times with at least 250 mL of DI water. Place mesh sieves of desired pore sizes (e.g., 50 &#181;m, 100 &#181;m, 300 &#181;m, 500 &#181;m) into each union joint with pore size decreasing from the top to the bottom of the filtration device (Figure 1A). Seal each union joint tightly to prevent leaking.</li>
	<li>Fold the mixed cellulose ester membrane filter(s) (142 mm diameter) of desired pore size(s) (e.g., 0.45 &#181;m) into a cone shape and place it into the filtration device: 
	Note: Folding the membrane filter will provide more surface area to prevent clogging of the filter.
	<ol>
		<li>Wet the membrane filter with DI water. While damp, fold the membrane filter into a cone shape with a diameter that fits into the filtration device. Also, fold a small lip along the edge of the cone so that it fits over the top of the union joint (Figure 1B). 
		Caution: The membrane filter must be wet before folding to prevent tearing.</li>
		<li>Place the stainless steel mesh basket into the last union joint (Figure 1C). Carefully place the cone-shaped membrane filter into the basket (Figure 1D). Fold the lip of the membrane filter over the edge of the union joint. 
		Note: The mesh basket will support the filter and reduce breakage once a vacuum has been applied.</li>
	</ol>
	</li>
	<li>Place a mesh sieve with the smallest desired pore size (e.g., 50 &#181;m) on top of the membrane filter in the last union joint seen in Figure 1. 
	Note: This will provide extra support to hold the membrane filter in place during filtration.</li>
	<li>Once all union joints are sealed tightly, attach the hose from the top of the filtering flask to the base of the filtration device. Then attach the hose from the side of the filtering flask to the vacuum pump as illustrated in Figure 2.</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57969/57969fig1.jpg" /> 
Figure 1: Assembly of the filtration device. (A) The filtration device is assembled by placing mesh sieves of desired pore size into the upper union joints. (B) The mixed cellulose ester membrane filter(s) must be folded into a cone-shaped to fit the diameter of the filtration device; the cone should include a small lip to fit over the edge of the union joint to secure the filter in place. (C) A mesh basket is placed into the union to add stability to the membrane filter. (D) The folded membrane filter is added to the mesh basket and the smallest mesh sieve size is placed over the top of the membrane filter. (E) The fully assembled filtration device. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57969/57969fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57969/57969fig2.jpg" /> 
Figure 2: Assembly of the filtering flask and pump. A filtering flask is attached to the filtration device vacuum adapter using a clear vinyl tubing. The filtering flask is then attached to the vacuum pump. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/57969/57969fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Sample Filtration
<ol>
	<li>Collect equipment blanks prior to filtration each time the device is assembled. Thoroughly rinse the device three times with at least 250 mL of DI water before the blank is collected. These blanks are collected using the steps outlined in steps 4.2 - 4.4.</li>
	<li>Turn on the vacuum pump. Ensure that the pressure of the vacuum pump does not exceed 127 mm Hg, or the membrane filter could tear. 
	Caution: Depending on the flow rate of sample filtration, pressure could increase inside the filtration device if sediment clogs the mesh sieves or membrane filters. This could potentially lead to a rupture in the membrane filter before reaching a reading of 127 mm Hg. For this reason, watch the pressure closely as it may need to be adjusted below 127 mm Hg on a sample by sample basis.</li>
	<li>Use a 500 mL graduated cylinder, triple rinsed with at least 250 mL of DI water, to measure the total volume of the sample. Record the volume and transfer the sample from the graduated cylinder to the filtration device. 
	Caution: Depending on the size of the water sample and the filtering flask, the filtering flask may need to be emptied multiple times during sample filtration.
	<ol>
		<li>To empty the filtering flask, turn off the pump and detach the two hoses from the flask. Empty the flask into a separate waste container. 
		Caution: Keep the filtered sample water until the entire sample has been filtered and it is confirmed that the membrane filter is intact.</li>
		<li>To continue the filtration cycle, reattach the hoses to the filtering flask, as outlined in step 3.4, and turn on the pump.</li>
	</ol>
	</li>
	<li>Once the entire sample has been filtered, rinse the sample container and graduated cylinder three times with at least 250 mL of DI water. After each rinse, filter the water used to rinse the container and graduated cylinder to ensure all particulates have been filtered.</li>
</ol>
5. Microplastic Filtration Device Disassembly
<ol>
	<li>Rinse the walls of the filtration device three times with at least 250 mL of DI water to ensure that all particulates have been filtered and none remain on the filtration device.</li>
	<li>Turn off the vacuum pump, then carefully unscrew and detach the first union. Turn the pump back on and use a DI water wash bottle to rinse the edges of the union joint. Wash particulates at the edges of the mesh sieve into the center to ensure that they are all collected.</li>
	<li>Turn the pump off and remove the mesh sieve carefully with clean forceps, making sure not to touch the particulates on the surface of the mesh sieve. Place the mesh sieve into a covered Petri dish and dry it at 60 &#7506;C for 24 h. Once dry, samples can be stored until analysis can begin.</li>
	<li>Repeat steps 5.1 - 5.3 for each union joint housing a mesh sieve.</li>
	<li>For the last union joint that houses a mesh sieve and membrane filter, repeat steps 5.1-5.3 for the mesh sieve. 
	Caution: Be careful when rinsing the mesh sieve, as sample can be lost if rinsed under the membrane filter.</li>
	<li>Turn the vacuum pump on and rinse the edges of the membrane filter using a DI water wash bottle. Wash particulates at the edges of the membrane filter into the center to ensure the full sample is filtered. Before removing the membrane filter, ensure that all water has passed through it and that no water is pooling on its surface. 
	Caution: Again, be careful when rinsing the membrane filter as sample can be lost if rinsed under it.</li>
	<li>Carefully remove and unfold the membrane filter with the forceps. Place the membrane filter into a Petri dish or foil envelope appropriate for its diameter. 
	Note: The membrane filter must be damp while being handled to prevent tearing.</li>
	<li>Dry the covered membrane filter in the oven at 60 &#7506;C for 24 h. Once dry, store samples until analysis can begin. 
	Note: The protocol can be paused here.</li>
</ol>
6. Particulate Analysis
<ol>
	<li>Leave the mesh sieve or membrane filter in the Petri dish and remove only the lid to begin examining the sample for microplastics. This will ensure that if any particulates fall off the mesh sieve or membrane filter they will remain in the Petri dish, which can be analyzed after all particulates are removed from the mesh sieve or membrane filter.</li>
	<li>Examine the mesh sieve or membrane filter under a stereomicroscope (14 - 90X magnification) to identify suspected plastic particulates and fibers. Use the following criteria when identifying suspected plastics: no cellular structure, fibers are equal thickness throughout, and particles are not shiny24.</li>
	<li>Remove all suspected plastics from the mesh sieve or membrane filter and place them into a collection vial containing 70% ethanol. Record the color and shape (e.g., particulate, fiber, film, etc.) of each suspected plastic.</li>
	<li>Once all suspected plastics are removed from the mesh sieve or membrane filter and quantified, examine both the lid and bottom of the Petri dish following steps 6.2 - 6.3.</li>
	<li>After the mesh sieve or membrane filter and Petri dish have been examined and all suspected plastics removed and quantified, place the particulates or fibers from the collection vial onto a 12-slot aluminum coated slide for analysis using a micro ATR-FTIR. 
	Note: It is not always feasible to test every suspected plastic on the micro ATR-FTIR. Therefore, &#34;strategically choose&#34; the amount that will address the goals of the study and anomalies in the suspected plastics (e.g., a high number of similar fibers or particles)25. In a general sense, test as many suspected plastics as possible, but no less than 20%.
	<ol>
		<li>Once suspected plastics are analyzed using micro ATR-FTIR, use spectral databases to determine if a given sample is plastic and, if so, determine the plastic&#39;s polymer type.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Microplastic collection using drift nets is the conventional method in environments like the ocean where both sediment and plastic concentrations are low, thus requiring large sample volumes. However, drift nets are not always practical or safe in rivers with high sediment loads and large floating or submerged debris. Additionally, it is not feasible to use a drift net when attempting to thoroughly capture and quantify microplastic materials, particularly fibers, as most nets used for plastic surveys have mesh sizes &#88...

Plastic was first observed in the ocean as early as the 1930s1. Recent estimates of marine plastic debris range from over 243,000 metric tons (MT) of plastic on the ocean&#39;s surface to 4.8-12.7 million MT of plastic entering the ocean from terrestrial sources annually2,3. Early studies on marine plastic debris focused on macroplastics (&#62;5 mm diameter) as they are easily visible and quantifiable. However, it was recently discovered that macroplastics represent &#60;10% of plastic debris, by count, in the ocean, indicating that the overwhelming majority of plastic debris is microplastic (&#60;5 mm diameter)2.
Microplastics are categorized into two groups: primary and secondary microplastics. Primary microplastics consist of plastics that are manufactured at a diameter &#60;5 mm and include nurdles, the raw pellets used to make consumer products, microbeads used as exfoliants in personal care products (e.g., facial wash, body scrub, toothpaste), and abrasives or lubricants in industry. Secondary microplastics are created within the environment as larger plastic debris is fragmented by photolysis, abrasion, and microbial decomposition4,5. Synthetic fibers are also secondary microplastics and are a growing concern. A single garment can release &#62;1,900 fibers per wash in a domestic washing machine6. These microfibers, as well as microbeads from personal care products, are washed down drains and into the sewer system before entering wastewater treatments plants. Murphy (2016) found that a wastewater treatment plant serving a population of 650,000 reduced the microplastic concentration by 98.4% from influent to effluent, yet 65 million microplastics remained in effluent and sludge each day7. Even with high percentages of microplastics being removed during the treatment processes, millions, possibly billions, of microplastics pass through wastewater treatment plants daily and enter surface waters in effluent6,8,9,10,11.
Due to their environmental release, microplastics have been found in the digestive and respiratory tissues of marine organisms across all trophic levels12,13,14,15. Their impact after uptake is variable, with some studies not observing harm, while others demonstrate numerous effects such as physical and chemical tissue damage4,6,14,15. Due to these discoveries, interest in this field has increased over the past five decades. However, only recently have studies begun to quantify plastic debris, particularly microplastics, in freshwater systems, such as rivers and lakes, or assess the effect on organisms dwelling in these habitats12,16,17,18. Rivers are a major source of plastic debris found in the ocean as they receive wastewater effluent and surface water runoff that contain microplastics and macroplastics.
The protocol detailed here can be used to collect microplastic samples where drift nets are not feasible; specifically, in aquatic environments with high concentrations of suspended sediments and large floating debris like the Mississippi River. The Mississippi River watershed is one of the world&#39;s largest and has a population of &#62;90 million people, likely making it one of the largest sources of plastic debris to the ocean19,20. Each year, the Mississippi River discharges an average of 735 km3 of freshwater into the Gulf of Mexico, along with high concentrations of suspended sediments (~60 to &#62;800 mg/L) and large debris13,21. Water samples were collected at two depths (i.e., surface and 0.6-depth) at various locations along the Mississippi River and its tributaries in translucent 1 L low-density polyethylene (PE) containers using a peristaltic pump. In the lab, samples were filtered using nylon mesh sieves and mixed cellulose ester membrane filters simultaneously with a custom-made 63.5 mm (2.5 in) polyvinyl chloride (PVC) cylinder with union joints to insert the sieves and filters22. The inclusion of PVC unions in the filtration device allows for filtration by as many or as few particle size classes as desired. Additionally, it can be used to capture microplastic debris down to sub-micron sizes using membrane filters when studying synthetic fibers. Once filtered, samples were dried and suspected plastics were identified and sorted from the mesh sieves and membrane filters under a stereomicroscope. Suspected plastics were then examined using micro-attenuated total reflectance Fourier transform infrared spectroscopy (micro ATR-FTIR) to eliminate non-synthetic materials or determine polymer type. Considering the size of microplastic particulates and fibers, contamination is commonplace. Sources of contamination include atmospheric deposition, clothing, field and lab equipment, as well as deionized (DI) water sources. Multiple steps are included throughout the protocol to reduce contamination from various sources while conducting all stages of the study.

<table><tbody><tr><td>1L Cubitainer Containers, Low-Density Polyethylene</td><td>VWR</td><td>89094-140</td><td>Containers used to collect and store samples.</td></tr><tr><td>2-1/2&quot; Clear Schedule 40 Rigid PVC Pipe</td><td>United States Plastic Corporation</td><td>34138</td><td>The PVC pipe used to make the device comes as an 2.43 m pipe. The pipe was then cut to the desired lengths for each section seperated by union joints. Section lengths were decided by predicting smaller pore sizes would clogg the device quicker. Longer sections were placed above the smaller pore sizes to collect and hold water to prevent needing to disassemble the device to change a filter while a sample remained in the device. For one filtration device one 18 in, one 12 in, and two 6 in peices are needed.</td></tr><tr><td>2-1/2&quot; PVC SCH 40 Socket Union&amp;nbsp;</td><td>Supply House</td><td>457-025</td><td>Union joints were glued to PVC pipe to house nylon sieves and mixed cellulose membranes.</td></tr><tr><td>Nylon 6 Woven Mesh Sheet, Opaque Off-White, 12&quot; Width, 12&quot; Length, 500 microns Mesh Size, 38% Open Area (Pack of 5)</td><td>Small Parts via Amazon</td><td>CMN-0500-C/5PK-05</td><td>Mesh sheets were cut into circles to match the diameter of the outer diameter of the PVC pipe. The edges were glued to esure no fraying would occur. The glue &#039;s diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.&amp;nbsp;</td></tr><tr><td>Nylon 6 Woven Mesh Sheet, Opaque White, 12&quot; Width, 12&quot; Length, 100 microns Mesh Size, 44% Open Area (Pack of 5)</td><td>Small Parts via Amazon</td><td>B0043D1TB4</td><td>Mesh sheets were cut into circles to match the diameter of the outer diameter of the PVC pipe. The edges were glued to esure no fraying would occur. The glue &#039;s diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.&amp;nbsp;</td></tr><tr><td>Nylon 6 Woven Mesh Sheet, Opaque White, 12&quot; Width, 12&quot; Length, 50 microns Mesh Size, 37% Open Area (Pack of 5)</td><td>Small Parts via Amazon</td><td>B0043D1SGA</td><td>Mesh sheets were cut into circles to match the diameter of the outer diameter of the PVC pipe. The edges were glued to esure no fraying would occur. The glue &#039;s diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.&amp;nbsp;</td></tr><tr><td>Mixed Cellulose Ester Membrane, 0.45um, 142mm, 25/pk</td><td>VWR</td><td>10034-914</td><td>Mixed cellulose membrane filter with 0.45 um was used as the last filter. A large diameter was used to allow the filter to be folded into a cone to increase surface area of the filter to prevent clogging.&amp;nbsp;</td></tr><tr><td>Metal Mesh Basket Tea Leaves Strainer Teapot Filter 76mm Dia 3pcs</td><td>Uxcell via Amazon</td><td>a15071600ux0260</td><td>The mesh basket used to provide extra support for the membrane filter to prevent tearing when pressure was applied by a vacuum pump.</td></tr><tr><td>1/2&quot; PVC Barbed Insert Male Adapter</td><td>Supply House</td><td>1436-005</td><td>A vacuum adapter was added to allow vacuum filtration in the case of slow filtration due to high sediment concentration.</td></tr><tr><td>1/2 in. O.D. x 3/8 in. I.D. x 10 ft. PVC Clear Vinyl Tube</td><td>Home Depot</td><td>702229</td><td>Tubing used to connect the vacuum pump to the filtration device.</td></tr><tr><td>YSI Professional Plus Multiparameter Instrument with Quatro Cable</td><td>YSI</td><td>6050000</td><td>Handheld meter used to measure additional water quality parameters parameters (e.g., turbidity, temperature, conductivity, pH, and dissolved oxygen (DO)).</td></tr><tr><td>2100P Portable Turbidimeter</td><td>Hach</td><td>4650000</td><td>Handheld meter used to measure turbidity.</td></tr><tr><td>FEP-lined PE tubing</td><td>Geotech</td><td>87050529</td><td>Tubing used with perestaltic pump to collect water samples from desired depths.</td></tr><tr><td>Geopump Peristaltic Pump Series II</td><td>Geotech</td><td>91350123</td><td>Pump used to collected water samples.</td></tr><tr><td>MeiJi Techno EMZ-8TR Microscope</td><td>Microscope.com</td><td>EMZ8TR-PLS2</td><td>Microscope used analyze mesh sieves and membrane filters to quanitfy suspect microsplastics.</td></tr><tr><td>Nicolet iS10 FTIR Spectrometer</td><td>Thermo Electron North America</td><td>912A0607</td><td>FTIR used to analyze suspect microplastics.</td></tr><tr><td>Nicolet iN5 FTIR microscope</td><td>Thermo Electron North America</td><td>912A0895</td><td>FTIR microscope used to analyze suspect microplastics.</td></tr><tr><td>Germanium (Ge) ATR</td><td>Thermo Electron North America</td><td>869-174400</td><td>Geranium ATR accessory used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.</td></tr><tr><td>Aluminum EZ-Spot Micro Mounts (Pkg of 5)</td><td>Thermo Electron North America</td><td>0042-545</td><td>Microscope slides used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.</td></tr><tr><td>Aluminum Coated Glass Sample Slides</td><td>Thermo Electron North America</td><td>0042-544</td><td>Microscope slides used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.</td></tr></tbody></table>

sampling, sorting, and characterizing microplastics in aquatic environments with high suspended sediment loads and large floating debris

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only recently have microplastics in freshwater systems, such as rivers and lakes, been quantified. Microplastic sampling at the surface usually consists of deploying drift nets behind either a stationary or moving boat, which limits the sampling to environments with low levels of suspended sediments and floating or submerged debris. Previous studies that employed drift nets to collect microplastic debris typically used nets with &#8805;300 &#181;m mesh size, allowing plastic debris (particles and fibers) below this size to pass through the net and elude quantification. The protocol detailed here enables: 1) sample collection in environments with high suspended loads and floating or submerged debris and 2) the capture and quantification of microplastic particles and fibers &lt;300 &#181;m. Water samples were collected using a peristaltic pump in low-density polyethylene (PE) containers to be stored before filtering and analysis in the lab. Filtration was done with a custom-made microplastic filtration device containing detachable union joints that housed nylon mesh sieves and mixed cellulose ester membrane filters. Mesh sieves and membrane filters were examined with a stereomicroscope to quantify and separate microplastic particulates and fibers. These materials were then examined using a micro-attenuated total reflectance Fourier transform infrared spectrometer (micro ATR-FTIR) to determine microplastic polymer type. Recovery was measured by spiking samples using blue PE particulates and green nylon fibers; percent recovery was determined to be 100% for particulates and 92% for fibers. This protocol will guide similar studies on microplastics in high velocity rivers with high concentrations of sediment. With simple modifications to the peristaltic pump and filtration device, users can collect and analyze various sample volumes and particulate sizes.

To validate the recovery rates of this protocol, three samples (V1-V3) from Oso Bay, Corpus Christi, Texas (adjacent to the Texas A&#38;M University Corpus Christi Campus), were spiked with 10 blue PE particulates (ranging from 50-100 &#181;m in diameter) and 50 green nylon fibers of various lengths (Figure 3). Sample TSS was calculated (Section 2) and then the samples were filtered using the methods outlined in Sections 3-5. The blue PE...

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Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris

Most microplastic research to date has occurred in marine systems where suspended solid levels are relatively low. Focus is now shifting to freshwater systems, which may feature high sediment loads and floating debris. This protocol addresses collecting and analyzing microplastic samples from aquatic environments that contain high suspended solid loads.

The ubiquitous presence of plastic debris in the ocean is widely recognized by the public, scientific communities, and government agencies. However, only ...

sampling-sorting-characterizing-microplastics-aquatic-environments

Universidad San Sebastian

Research

JoVE Journal

Environment

15.9K Views.  Texas A&M University-Corpus Christi. Most microplastic research to date has occurred in marine systems where suspended solid levels are relatively low. Focus is now shifting to freshwater systems, which may feature high sediment loads and floating debris. This protocol addresses collecting and analyzing microplastic samples from aquatic environments that contain high suspended solid loads. 

Video: Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

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

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

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

Characterization and Application of Passive Samplers for Monitoring of Pesticides in Water

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study provides a protocol for the passive sampler preparation, calibration, extraction method and instrumental analysis. Sampling rates (RS) and passive sampler-water partition coefficients (KPW) were calculated for silicone rubber, polar organic chemical integrative sampler POCIS-A, POCIS-B, SDB-RPS and C18 disk. The uptake of the selected compounds depended on their physicochemical properties, i.e., silicone rubber showed a better uptake for more hydrophobic compounds (log octanol-water partition coefficient (KOW) &#62; 5.3), whereas POCIS-A, POCIS-B and SDB-RPS disk were more suitable for hydrophilic compounds (log KOW &#60; 0.70).

A protocol about the characterization and application of five different passive sampling devices is presented.

Five different water passive samplers were calibrated under laboratory conditions for measurement of 124 legacy and current used pesticides. This study ...

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

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

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

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

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

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 ...

Extraction of Organochlorine Pesticides from Plastic Pellets and Plastic Type Analysis

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment during manufacturing and transport. Because of their environmental persistence, they are widely distributed in the oceans and on beaches all over the world. They can act as a vector of potentially toxic organic compounds (e.g., polychlorinated biphenyls) and might consequently negatively affect marine organisms. Their possible impacts along the food chain are not yet well understood. In order to assess the hazards associated with the occurrence of plastic pellets in the marine environment, it is necessary to develop methodologies that allow for rapid determination of associated organic contaminant levels. The present protocol describes the different steps required for sampling resin pellets, analyzing adsorbed organochlorine pesticides (OCPs) and identifying the plastic type. The focus is on the extraction of OCPs from plastic pellets by means of a pressurized fluid extractor (PFE) and on the polymer chemical analysis applying Fourier Transform-InfraRed (FT-IR) spectroscopy. The developed methodology focuses on 11 OCPs and related compounds, including dichlorodiphenyltrichloroethane (DDT) and its two main metabolites, lindane and two production isomers, as well as the two biologically active isomers of technical endosulfan. This protocol constitutes a simple and rapid alternative to existing methodology for evaluating the concentration of organic contaminants adsorbed on plastic pieces.

Microplastics act as vector of potentially toxic organic contaminants with unpredictable effects. This protocol describes an alternative methodology for assessing the levels of organochlorine pesticides adsorbed on plastic pellets and identifying the polymer chemical structure. The focus is on pressurized fluid extraction and attenuated total reflectance Fourier transform infrared spectroscopy.

Plastic resin pellets, categorized as microplastics (&#8804;5 mm in diameter), are small granules that can be unintentionally released to the environment ...

An Ultra-clean Multilayer Apparatus for Collecting Size Fractionated Marine Plankton and Suspended Particles

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those of suspended/sinking particles. Here, we present an all plastic (Polypropylene and Polycarbonate), multi-layer filtration system for collection of suspended particulate matter (SPM) at sea. This ultra-clean sampling device has been designed and developed specifically for trace element studies. Meticulous selection of all non-metallic materials and utilization of an in-line flow-through procedure minimizes any possible metal contamination during sampling. This system has been successfully tested and tweaked for determining trace metals (e.g., Fe, Al, Mn, Cd, Cu, Ni) on particles of varying size in coastal and open ocean waters. Results from the South China Sea at the South East Asia Time-Series (SEATS) station indicate that diurnal variations and spatial distribution of plankton in the euphotic zone can be easily resolved and recognized. Chemical analysis of size-fractionated particles in surface waters of the Taiwan Strait suggests that the larger particles (&#62;153 &#181;m) were mostly biologically derived, while the smaller particles (10 - 63 &#181;m) were mostly composed of inorganic matter. Apart from Cd, the concentrations of metals (Fe, Al, Mn, Cu, Ni) decreased with increasing size.

Plankton and suspended particles play a major role in the biogeochemical cycles in the ocean. Here, we provide an ultra-clean, low stress method for the collection of various sizes of particles and plankton at sea with the capability of handling large volumes of seawater.

The distributions of many trace elements in the ocean are strongly associated with the growth, death, and re-mineralization of marine plankton and those ...

Characterization of Aquatic Biofilms with Flow Cytometry

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond quickly to environmental changes and can be thus used as indicators of water quality. Currently, biofilm assessment is mostly based on integrative and functional endpoints, such as photosynthetic or respiratory activity, which do not provide information on the biofilm community structure. Flow cytometry and computational visualization offer an alternative, sensitive, and easy-to-use method for assessment of the community composition, particularly of the photoautotrophic part of freshwater biofilms. It requires only basic sample preparation, after which the entire sample is run through the flow cytometer. The single-cell optical and fluorescent information is used for computational visualization and biological interpretation. Its main advantages over other methods are the speed of analysis and the high-information-content nature. Flow cytometry provides information on several cellular and biofilm traits in a single measurement: particle size, density, pigment content, abiotic content in the biofilm, and coarse taxonomic information. However, it does not provide information on biofilm composition on the species level. We see high potential in the use of the method for environmental monitoring of aquatic ecosystems and as an initial biofilm evaluation step that informs downstream detailed investigations by complementary and more detailed methods.

Flow cytometry in combination with visual clustering offers an easy-to-use and fast method for studying aquatic biofilms. It can be used for biofilm characterization, detection of changes in biofilm community structure, and detection of abiotic particles embedded in the biofilm.

Biofilms are dynamic consortia of microorganism that play a key role in freshwater ecosystems. By changing their community structure, biofilms respond ...

Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish

As a new type of environmental pollutant, microplastic has been widely found in the aquatic environment and poses a high threat to aquatic organisms. The bioaccumulation of microplastics plays a key role in their toxic effects; however, as a particulate, their bioaccumulations are different from many other pollutants. Described here is a feasible method to visually determine the accumulation and distribution of microplastics in zebrafish embryos or larvae using fluorescent microplastics. Embryos are exposed to different concentrations (0.1, 1, and 10 mg/L) of fluorescent microplastics with a diameter of 500 nm for 120 h. It is shown in the results that microplastics can bioaccumulate in zebrafish embryos/larvae in a concentration-dependent manner. Before hatching, strong fluorescence is found around the embryonic chorion; while in zebrafish larvae, the yolk sac, pericardium, and gastrointestinal tract are the main accumulated sites of microplastics. The results demonstrate the uptake and internalization of microplastics in zebrafish at early life stages, which will provide basis for better understanding the impact of microplastics on aquatic animals.

Zebrafish embryos/larvae develop externally and are optically transparent. The bioaccumulation of microplastics in fish at early life stages is readily assessed with fluorescently labeled microbeads.

As a new type of environmental pollutant, microplastic has been widely found in the aquatic environment and poses a high threat to aquatic organisms. The ...

Sampling, Identification and Characterization of Microplastics Release from Polypropylene Baby Feeding Bottle during Daily Use

Microplastics (MPs) are becoming a global concern due to the potential risk to human health. Case studies of plastic products (i.e., plastic single-use cups and kettles) indicate that MP release during daily use can be extremely high. Precisely determining the MP release level is a crucial step to identify and quantify the exposure source and assess/control the corresponding risks stemming from this exposure. Though protocols for measuring MP levels in marine or freshwater has been well developed, the conditions experienced by household plastic products can vary widely. Many plastic products are exposed to frequent high temperatures (up to 100 &#176;C) and are cooled back to room temperature during daily use. It is therefore crucial to develop a sampling protocol that mimics the actual daily-use scenario for each particular product. This study focused on widely used polypropylene-based baby feeding bottles to develop a cost-effective protocol for MP release studies of many plastic products. The protocol developed here enables: 1) prevention of the potential contamination during sampling and detection; 2) realistic implementation of daily-use scenarios and accurate collection of the MPs released from baby feeding bottles based on WHO guidelines; and 3) cost-effective chemical determination and physical topography mapping of MPs released from baby feeding bottles. Based on this protocol, the recovery percentage using standard polystyrene MP (diameter of 2 &#181;m) was 92.4-101.2% while the detected size was around 102.2% of the designed size. The protocol detailed here provides a reliable and cost-effective method for MP sample preparation and detection, which can substantially benefit future studies of MP release from plastic products.

This study detailed a reliable and cost-effective protocol for microplastics collection and detection from the daily use of plastic products.

Microplastics (MPs) are becoming a global concern due to the potential risk to human health. Case studies of plastic products (i.e., plastic single-use ...

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.

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.

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

Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris

Summary

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Chapters in this video

Sediment Core Sectioning and Extraction of Pore Waters under Anoxic Conditions

Characterization and Application of Passive Samplers for Monitoring of Pesticides in Water

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

Protocol for Microplastics Sampling on the Sea Surface and Sample Analysis

Extraction of Organochlorine Pesticides from Plastic Pellets and Plastic Type Analysis

An Ultra-clean Multilayer Apparatus for Collecting Size Fractionated Marine Plankton and Suspended Particles

Characterization of Aquatic Biofilms with Flow Cytometry

Accumulation and Distribution of Fluorescent Microplastics in the Early Life Stages of Zebrafish

Sampling, Identification and Characterization of Microplastics Release from Polypropylene Baby Feeding Bottle during Daily Use

Separation and Identification of Conventional Microplastics from Farmland Soils