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

Katherine M. Martin; Elizabeth A. Hasenmueller; John R. White; Lisa G. Chambers; Jeremy L. Conkle

doi:10.3791/57969

JoVE Journal > Environment

Environnement

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

Published: July 28, 2018

doi:

10.3791/57969

Katherine M. Martin¹, Elizabeth A. Hasenmueller², John R. White³, Lisa G. Chambers⁴, Jeremy L. Conkle¹

¹Department of Physical and Environmental Sciences,Texas A&M University-Corpus Christi, ²Department of Earth and Atmospheric Sciences,Saint Louis University, ³Department of Oceanography and Coastal Sciences,Louisiana State University, ⁴Department of Biology,University of Central Florida

Summary

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.

Abstract

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 ≥300 µm 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 <300 µm. 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.

Introduction

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'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 (>5 mm diameter) as they are easily visible and quantifiable. However, it was recently discovered that macroplastics represent <10% of plastic debris, by count, in the ocean, indicating that the overwhelming majority of plastic debris is microplastic (<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 <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 >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's largest and has a population of >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 >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.

Protocol

1. Water Sample Collection

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.
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).
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.
1. 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.
Place the effluent end of the tubing over the boat'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.
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).
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.
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.
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.
Store the microplastic and TSS subsamples on ice until returning to the lab, where they will be stored at -20°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.

2. TSS Determination

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.

3. Microplastic Filtration Device Assembly

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 µm, 100 µm, 300 µm, 500 µm) 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.
Fold the mixed cellulose ester membrane filter(s) (142 mm diameter) of desired pore size(s) (e.g., 0.45 µm) 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.
1. 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.
2. 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.
Place a mesh sieve with the smallest desired pore size (e.g., 50 µm) 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.
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.

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. Please click here to view a larger version of this figure.

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. Please click here to view a larger version of this figure.

4. Sample Filtration

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.
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.
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.
1. 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.
2. To continue the filtration cycle, reattach the hoses to the filtering flask, as outlined in step 3.4, and turn on the pump.
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.

5. Microplastic Filtration Device Disassembly

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.
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.
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 ᵒC for 24 h. Once dry, samples can be stored until analysis can begin.
Repeat steps 5.1 – 5.3 for each union joint housing a mesh sieve.
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.
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.
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.
Dry the covered membrane filter in the oven at 60 ᵒC for 24 h. Once dry, store samples until analysis can begin.
Note: The protocol can be paused here.

6. Particulate Analysis

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.
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.
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.
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.
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, "strategically choose" 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%.
1. 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's polymer type.

Representative Results

To validate the recovery rates of this protocol, three samples (V₁-V₃) from Oso Bay, Corpus Christi, Texas (adjacent to the Texas A&M University Corpus Christi Campus), were spiked with 10 blue PE particulates (ranging from 50-100 µm 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 particulates and green nylon fibers were then separated and quantified (Table 1). Other fibers and particulates were observed on the mesh sieves and membrane filters, likely derived from the Oso Bay water sample. On average, 100% of the PE particulates and 92% of the nylon fibers were recovered. A loss of fibers may be due to a small amount of sample loss during filtration or incorrect identification.

An equipment blank was collected from the filtration device by filtering 1000 mL of DI water. This blank was analyzed using 100 µm and 50 µm mesh sieves and a 0.45 µm membrane filter. A total of 7 fibers (blue and clear) were found in the equipment blank. This contamination could have been from the filtration device, laboratory equipment, atmospheric deposition, or DI water. However, the fibers were not similar to the blue PE particulates and green nylon fibers used to spike the samples.

This protocol was created to process samples from the Mississippi River watershed, including the Mississippi River mainstem and the Missouri River. Preliminary analyses from the Mississippi River and Missouri River had an average TSS of 63 mg/L. While the TSS values of Oso Bay are typically below those observed in the Mississippi River watershed, sediment was intentionally disturbed prior to water collection to simulate higher suspended sediment concentrations that might be encountered in large river systems. The average TSS in the Oso Bay samples was 1,865 mg/L, which is ~30 times higher than the TSS calculated for the Mississippi River and Missouri River samples. The turbid Oso Bay samples suggest successful filtration for samples with a TSS of up to ~1,800 mg/L using the techniques outlined here.

Figure 3: Particulates and fibers used for percent recovery validation. Image of two blue PE particulates and two green nylon fibers in a range of sizes used to spike the validation samples from Oso Bay in Corpus Christi, Texas. Please click here to view a larger version of this figure.

Sample	TSS (g/L)	0.45 μm		50 μm		100 μm		Total		% Recovered
		Fibers	Particulates	Fibers	Particulates	Fibers	Particulates	Fibers	Particulates	Fibers	Particulates
V1	4.663	1	0	18	0	31	10	50	10	100	100
V2		0	0	21	0	28	10	49	10	98	100
V3		0	0	27	0	14	10	41	10	82	100

Table 1: Results from validation samples. A set number of blue PE particulates and green nylon fibers were added to samples taken from Oso Bay in Corpus Christi, Texas, to validate the filtration device and analysis protocol. Three microplastic validation samples (V₁-V₃) and one TSS sample were taken at the same location at the bank of Oso Bay. The fibers and particulates were quantified for each pore size and a total was calculated for each validation sample. Using the known amount of fibers and particulates used to spike the samples and the total recovered from each sample, the percent recovery was calculated.

The protocol was also designed to sample rivers from two depths: the surface (the river depth with the highest velocity) and 0.6-depth (the river depth with approximately average velocity for the entire water column). Samples from the Mississippi River and Missouri River (Table 2) were collected and analyzed as described above. To examine the effect of depth on microplastic concentration, the first and second samples were taken at the same location (i.e., the Mississippi River at Alton, Illinois) but at different depths. To examine the possible effect of sampling location on microplastic loading, the first and third samples were taken at the same depth but at different locations (i.e., the Mississippi River at Alton, Illinois, and the Missouri River above Saint Louis, Missouri). Examples of the fibers and particulates found in the preliminary Mississippi River basin samples are shown in Figure 4.

Location	USGS Gauging Station	Depth	Turbidity	TSS	Fibers	Particles	Fibers	Particles	Fibers	Particles	Fibers	Particles	Total	Fiber/ Particule Ratio
		Depth	Turbidity	TSS	0.45 mm		50 mm		100 mm		Total		Total
		m	NTU	g/L	#/L
MS; Alton, IL	USGS 05587498	0	38.3	0.063	80	0	126	1	54	1	260	2	262	130
MS; Alton, IL	USGS 05587498	20.1	61.4	0.090	191	0	151	5	195	1	537	6	543	90
MO; Columbia Bottom, MO	USGS 06935965	0	30.8	0.036	122	4	57	0	37	0	216	4	220	54
MS = Mississippi River; MO = Missouri River

Table 2: Mississippi River watershed sample collection and analysis data. Preliminary samples were collected near USGS gauging stations at the Mississippi River and Missouri River. Depth (m), turbidity (NTU), and TSS (mg/L) were measured for each site. Samples were filtered and analyzed following this protocol. Fibers and particulates were quantified for 50 µm and 100 µm pore size mesh sieves as well as a 0.45 µm membrane filter. Due to a lack of materials collected on a 500 µm mesh sieve, this size is excluded from the results presented.

Figure 4: Example particulates and fibers found in preliminary samples from the Mississippi River watershed. Images of fibers and particulates quantified in a sample (Table 2) taken from the surface of the Mississippi River at Alton, Illinois. (A) Image of two blue fibers that range in size on a 0.45 µm membrane filter. (B) Image of a red particulate and various fibers found on a 50 µm mesh sieve, showing the range in color, size, and shape of the microplastics found in the Mississippi River watershed. Please click here to view a larger version of this figure.

Discussion

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 ≥300 µm. The protocol described in this paper allows for sampling in waterbodies containing high sediment loads while also permitting the capture of microplastics <300 µm in diameter. The method and associated filtering device are versatile and can be adapted to specific project needs. Furthermore, data obtained with this protocol will help develop mitigation strategies to improve water quality and measure the effectiveness of these strategies, such as the recent microbead ban²⁶.

This method enables control of sample collection depth, volume input, and separation of microplastics into size classes while accounting for multiple sources of contamination. Employing a peristaltic pump permits the user to collect samples at any desired depth by adjusting the length of the pump tubing. Users can easily control the sample volume with the use of the filtration device, while the detachable union fittings allow for adjustments in filter material and pore sizes to accommodate variable diameters and concentrations of plastic. We found that a 1 L sample size was ideal for quantifying microplastics in the Mississippi River watershed for several reasons. First, within 1 L of water, we found that there were several hundred suspected fibers and particles. Second, the high sediment masses in samples with volumes larger than 1 L slowed filtering substantially. Third, longer filtering times could potentially lead to greater lab contamination. The filtration device and the ability to easily adapt it to differing project needs facilitate the collection and analysis of microplastic debris at sub-micron sizes, which is particularly helpful when studying synthetic fibers.

The inclusion of union joints eases the removal of mesh sieves or membrane filters between filtration cycles but requires that joints be sealed firmly and carefully to ensure mesh sieves and membrane filters are seated properly, which prevents the loss of sample (Sections 3 and 5). To prevent tearing or cracking, the membrane filter needs to be damp before handling it, but dry before microscope analysis. Rupturing can occur in the membrane filter before the pump pressure reaches 127 mm Hg (steps 4.2), especially in samples with high sediment volume. Therefore, the pressure must be watched carefully and adjust as needed.

Though the protocol for using the filtration device alleviates problems associated with deploying drift nets such as clogging of the net with suspended sediments, it increases sample processing in the lab, which increases the chances for contamination. To reduce or eliminate potential contamination from sample handling, all equipment must be thoroughly rinsed with sufficient quantities of DI water three times and blanks must be taken from each device (e.g., peristaltic pump, filtration device, collection container) throughout sample collection, processing, and analysis. Each environment and equipment blank will then be filtered and analyzed using the protocol outlined in Sections 4-6. The use of an ultra-pure water filtration system could reduce potential contamination from DI water used for rinsing and blanks.

In the lab, at least 20% of the samples should be analyzed by two individuals to ensure consistent plastic identification. During filtration and analysis in the lab, open Petri dishes can serve as lab blanks and be placed in designated areas for the duration of the analysis period. Each lab blank will then be analyzed using the protocol in Section 6. To prevent contamination from atmospheric deposition, cover all equipment with aluminum foil after washing with DI water.

The use of a peristaltic pump and custom-made microplastic filtration device in this protocol allows users to collect samples in environments containing high concentrations of suspended sediments. Additionally, this method allows users to capture and quantify microplastic debris <300 µm, specifically microfibers. The percent recovery for this protocol was measured to be 100% and 92% for PE particulates and nylon fibers, respectively, showing relatively high recovery rates. Preliminary samples were taken in the Mississippi River watershed also using this protocol where 1 L samples averaged >200 microplastics ranging in size (0.45-500 µm), shape, and color. This protocol will guide similar studies on the fate, effects, and sources of microplastics.

Divulgations

The authors have nothing to disclose.

Acknowledgements

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.

Materials

1L Cubitainer Containers, Low-Density Polyethylene	VWR	89094-140	Containers used to collect and store samples.
2-1/2" Clear Schedule 40 Rigid PVC Pipe	United States Plastic Corporation	34138	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.
2-1/2" PVC SCH 40 Socket Union	Supply House	457-025	Union joints were glued to PVC pipe to house nylon sieves and mixed cellulose membranes.
Nylon 6 Woven Mesh Sheet, Opaque Off-White, 12" Width, 12" Length, 500 microns Mesh Size, 38% Open Area (Pack of 5)	Small Parts via Amazon	CMN-0500-C/5PK-05	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 's diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.
Nylon 6 Woven Mesh Sheet, Opaque White, 12" Width, 12" Length, 100 microns Mesh Size, 44% Open Area (Pack of 5)	Small Parts via Amazon	B0043D1TB4	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 's diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.
Nylon 6 Woven Mesh Sheet, Opaque White, 12" Width, 12" Length, 50 microns Mesh Size, 37% Open Area (Pack of 5)	Small Parts via Amazon	B0043D1SGA	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 's diamter should not extend into the inner diameter of the PVC so that it will not be affected during filtration.
Mixed Cellulose Ester Membrane, 0.45um, 142mm, 25/pk	VWR	10034-914	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.
Metal Mesh Basket Tea Leaves Strainer Teapot Filter 76mm Dia 3pcs	Uxcell via Amazon	a15071600ux0260	The mesh basket used to provide extra support for the membrane filter to prevent tearing when pressure was applied by a vacuum pump.
1/2" PVC Barbed Insert Male Adapter	Supply House	1436-005	A vacuum adapter was added to allow vacuum filtration in the case of slow filtration due to high sediment concentration.
1/2 in. O.D. x 3/8 in. I.D. x 10 ft. PVC Clear Vinyl Tube	Home Depot	702229	Tubing used to connect the vacuum pump to the filtration device.
YSI Professional Plus Multiparameter Instrument with Quatro Cable	YSI	6050000	Handheld meter used to measure additional water quality parameters parameters (e.g., turbidity, temperature, conductivity, pH, and dissolved oxygen (DO)).
2100P Portable Turbidimeter	Hach	4650000	Handheld meter used to measure turbidity.
FEP-lined PE tubing	Geotech	87050529	Tubing used with perestaltic pump to collect water samples from desired depths.
Geopump Peristaltic Pump Series II	Geotech	91350123	Pump used to collected water samples.
MeiJi Techno EMZ-8TR Microscope	Microscope.com	EMZ8TR-PLS2	Microscope used analyze mesh sieves and membrane filters to quanitfy suspect microsplastics.
Nicolet iS10 FTIR Spectrometer	Thermo Electron North America	912A0607	FTIR used to analyze suspect microplastics.
Nicolet iN5 FTIR microscope	Thermo Electron North America	912A0895	FTIR microscope used to analyze suspect microplastics.
Germanium (Ge) ATR	Thermo Electron North America	869-174400	Geranium ATR accessory used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.
Aluminum EZ-Spot Micro Mounts (Pkg of 5)	Thermo Electron North America	0042-545	Microscope slides used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.
Aluminum Coated Glass Sample Slides	Thermo Electron North America	0042-544	Microscope slides used along with the Nicolet iN5 FTIR microscope to analyze suspect microplastic.

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Martin, K. M., Hasenmueller, E. A., White, J. R., Chambers, L. G., Conkle, J. L. Sampling, Sorting, and Characterizing Microplastics in Aquatic Environments with High Suspended Sediment Loads and Large Floating Debris. J. Vis. Exp. (137), e57969, doi:10.3791/57969 (2018).