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Article

Microplastic and Nanoplastic Particle Isolation from Liquid and Biological Samples via Mini-Extruder Filtration (MEF)

by
Abigail Hall
1,
Luis F. Cardona Polo
1,
Kennedy Helms
1,
Alexei Christodoulides
1 and
Nathan J. Alves
1,2,*
1
Department of Emergency Medicine, Indiana University School of Medicine, Indianapolis, IN 46202, USA
2
Weldon School of Biomedical Engineering, Purdue University, West Lafayette, IN 47907, USA
*
Author to whom correspondence should be addressed.
Environments 2024, 11(8), 180; https://doi.org/10.3390/environments11080180
Submission received: 16 July 2024 / Revised: 11 August 2024 / Accepted: 16 August 2024 / Published: 21 August 2024
(This article belongs to the Special Issue Deployment of Green Technologies for Sustainable Environment III)
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Abstract

:
Microplastic pollution poses an increasing environmental and human health risk and additional techniques are needed to facilitate nondestructive, quantitative particle recovery and analysis. Using a mini-extruder filtration (MEF) device, the efficiency of pristine particle capture from solution and digested biological tissue (blood clots) was assessed. Polystyrene particles in both the submicron (100, 300, and 500 nm) and micron range (2, 5, 7, and 10 µm) with aminated, carboxylated, or unmodified surface modifications were explored. The MEF-isolated-particle recovery was analyzed pre- and postseparation isolation and quantified via a Nanosight LM10 particle tracking system (submicron particles) or hemacytometer (micron particles). Particles’ surface chemistry and concentration did not impact recovery compared to unfiltered samples with smaller particle sizes reducing recovery efficiency. Micron particle size recovery averaged 86.8 ± 4.3% across all surface chemistries at the same concentration; however, submicron particle recoveries varied by size and charge with 500 nm exhibiting recoveries of 80.6 ± 16.6%, 300 nm 73.0 ± 10.4%, and 100 nm particles 17.0 ± 10.3%. The mini-extruder device, used as a filtration recovery system, efficiently captures 10 to 0.5 µm particles from environmental and tissue samples making it an effective and low-cost platform facilitating the nondestructive capture of diverse microplastics for subsequent analysis.

Graphical Abstract

1. Introduction

In the 70 years since the widespread commercialization of plastics in the 1950s, plastic production has skyrocketed to 460 million tons per year in 2019 [1]. Plastic waste covers a diverse range of polymers, including polypropylene, high-density and low-density polyethylene, polystyrene, poly(vinyl chloride), and poly(ethylene terephthalate), to list a few [2,3]. This plastic debris is leaving a lasting legacy in the environment, resulting in damage to aquatic [4,5,6] and terrestrial ecosystems [7,8], and it is increasingly being identified as a possible danger to human health [9,10]. Microplastics are plastic particles or plastic debris smaller than 5 mm, and nanoplastics have several working definitions but are commonly defined as plastic particles between 1 and 1000 nm, often displaying colloidal characteristics [11]. Plastic has been quantified in many human tissues and biological fluids, including stool, lung, circulating blood and clots, tonsil tissue, and liver tissue [12,13,14,15,16,17]. The health effects of plastic exposure are a rapidly growing field of study, with both in vitro and in vivo models being investigated across wide ranges of particle sizes, shapes, and polymer types [18,19,20,21,22]. A major limitation of the current plastic particle research is the difficulty of working with smaller plastic particles, including <10 µm microplastics and nanoplastics [23,24].
Research has been conducted across a diverse range of particle sizes, with in vivo testing showing that plastics in the submicron-size range appear to be the most concerning from a physiological perspective and less understood than larger particles [25,26,27,28,29]. The quantification and recovery of intact particles from primary human and animal tissue samples are often more complicated, as particle sizes become smaller resulting in most studies not measuring particles <10–50 µm [30,31,32,33]. Plastic particle testing in animal models involve set treatments of pristine particles at defined concentrations, but due to difficulties associated with recovery and quantification, the total particle burden with absolute particle tracking of intake, excretion, and bio-burden is rarely carried out [34]. Current methods of quantifying and characterizing particles include Raman spectroscopy, dynamic light scattering, pyrolysis coupled with gas chromatography–mass spectrometry (Py-GC/MS), and flow cytometry [35,36,37]. Each of these methods comes with their own set of benefits and limitations providing either physical or chemical information on the nano-/microplastic sample. The methodologies used in sample preparation for these analysis techniques are also diverse, including various methods of preprocessing, such as ultracentrifugation, precipitation, selective staining, and filtration, with some techniques being destructive to the sample [38,39,40,41,42]. The field of plastic particle characterization continues to evolve as new environmental and health risks are identified.
Filtration is widely used to remove undesirable sample components during biological and environmental sample preparation or to capture particles for qualitative analysis. Because of common filtration issues, such as frequent membrane fouling and difficulty removing particles bound to membrane surfaces, quantitative capture of particles is rarely used for nanoplastics and smaller microplastics. To address the current limitations in nano-/microplastic isolation from diverse sample types, a simple and efficient use of a mini-extruder filtration (MEF) device is described herein. The mini-extruder discussed throughout this manuscript is designed for liposome formation applications and consists of two glass syringes that fit into a sealed stainless-steel chamber that holds a polycarbonate membrane filter with highly controlled pore sizes [43]. The inexpensive filters can be obtained in many pore sizes (0.03 to 12 µm), and the whole unit offers an affordable and reproducible nano-/microplastic isolation system. This accessible isolation technique lowers the barrier to entry for the recovery and quantification of micron and submicron particles from diverse biological and environmental liquid samples. To test the feasibility of using the MEF system for quantitative particle recovery, fluorescent polystyrene particles were obtained in a range of sizes and surface modifications, diluted into buffer, and filtered using the MEF system (see Figure 1A).
Additionally, recovery of microplastic-spiked blood clots formed under shear was also explored following potassium hydroxide (KOH) and enzymatic digestion as an additional proof-of-concept. Plastic-burdened blood clots were formed by clotting citrated human doner blood with a known concentration of 2, 5, 7, and 10 µm particles, and then the clots were digested and filtered using the MEF device to test the limits of the system with biological samples. The experiments presented herein test the viability and effective working parameters for using the MEF device to recover diverse particles (unfunctionalized, carboxylated, and aminated) across a wide range of particle sizes (100 nm–10 µm). This nondestructive workflow allows for more effective quantification of the total sample particle load, efficiently recovering particles for additional downstream particle analysis to be carried out including the following: particle count, particle surface characterization, particle shape analysis, and polymer material typing.

2. Materials and Methods

2.1. Materials and Particle Handling

Fluorescent polystyrene particles were purchased from MagSphere (Pasadena, CA, USA; 100 nm–10 µm) with unmodified, aminated, or carboxylated surface modifications for a total of 21 particle types (PSF-100NM, PSF-300NM, PSF-500NM, PSF-002UM, PSF-005UM, PSF-007UM, PSF-010UM, AMF-100NM, AMF-300NM, AMF-500NM, AMF-002UM, AMF-005UM, AMF-007UM, AMF-010UM, CMF-100NM, CMF-300NM, CMF-500NM, CMF-002UM, CMF-005UM, CMF-007UM, and CMF-010UM). Particle working stocks were prepared in 0.01% Tween 20 and 2 mg/mL bovine serum albumin (Sigma Aldrich, St. Louis, MO, USA; A7906), bath sonicated, and allowed to disperse overnight before testing. Each subsequent dilution was incubated >30 min before analysis. The mini-extruder (Avanti 610020), 1000 µL syringe (Avanti 610017), 10 mm filter supports (Avanti 610014), 19 mm 0.03 µm polycarbonate (PC) membranes (Avanti 61002), 19 mm 0.05 µm PC membranes (Avanti 61003-1Ea), and mini-extruder holding block (Avanti 610024) were purchased from Avanti Polar Lipids (Alabaster, AL, USA). When noted, heated reagents were brought to 55 °C in a Fisher Scientific Isotemp 105 heated water bath. Dilutions were performed in normal buffers prepared with deionized (DI) water as the use of fluorescent particles allowed for ignoring submicron particle laboratory contamination. If using nonfluorescent particles in the submicron range particle free water and additional particle negative controls should be used. Healthy human blood for research was obtained from consented doners under an approved IRB (protocol #1610652271) at Indiana University School of Medicine with blood samples being drawn by a trained phlebotomist.

2.2. Mini-Extruder Filtration (MEF)

Dilutions were prepared in 2 mg/mL bovine serum albumin (BSA) + 0.01% Tween 20 and sonicated utilizing a BRANSON ultrasonic water bath for 5 min and then allowed to disperse for at least 2 h. The MEF system was assembled using a 0.05 µm filter with only one filter support opposite the load syringe (see Figure 1B). The assembled system was prewet with a 1 mL flow of warm 55 °C DI water, using a 1000 µL glass loading syringe, through the filter to the flow through (FT) syringe, after which the FT was discarded. Then, the sample was loaded by flowing through the filter, manually applying consistent pressure on the syringe to maximize the particle capture [44,45]. After the sample loading, the system was washed with 1 mL of warm water and then 500 µL of air was pushed through to clear excess liquid from the main filter chamber. The chamber was then carefully disassembled, and the filter was removed and placed directly into a glass scintillation vial containing 400 µL of warm 0.01% Tween 20. The vial was sonicated in a bath sonicator for 3 min and then incubated on an orbital shaker for 10 min. To maximize particle recovery, the filter was exposed to an additional 400 µL (bringing the total vial volume to 800 µL), sonicated for 3 min, and incubated an additional 10 min (see Supplementary Protocol S1). After rinsing was completed, the filter was removed from the solution, and the vial was stored at room temperature in the dark until analysis.

2.3. Nanosight LM10 Analysis of Submicron Particles

The submicron particle size and recovery concentrations were analyzed on a Nanosight LM10 NTA (488 nmex/500 nm fluorescence filter) using the recommended settings from the manufacture for fluorescent particles. Briefly, the camera gain was set to 1500 for 500 and 300 nm particles and 800 for 100 nm particles, with 3 cycles of 1 min for data collection carried out on all samples pre- and post-MEF isolation. Each particle type and particle concentration condition was prepared in three separate sample replicates, each being run through the MEF device independently on the same day as the analysis, and measured with three technical replicates.

2.4. Hemacytometer and Light Microscope Analysis of Micron Particles

Micron-sized particle recovery concentrations were measured using a Hausser Bright-Line Improved Neubauer ruled hemacytometer (# 3110) on an upright light microscope (Leica CME Binocular Microscope; Deerfield, IL, USA) using the 40× objective lens. Each sample was loaded into both chambers of the hemacytometer, left to settle for 20 min, and then counted and averaged to determine the final particle concentration. Micron particle sizes were verified by brightfield imaging on a Nikon Eclipse Ni upright microscope, and image analysis was performed using QuPath v0.5.1 software. A minimum of 50 particle measurements were averaged for all micron-particle-sizing measurements.

2.5. Blood Clot Formation by Chander Loop

Healthy human blood was drawn into citrated blood collection tubes. A chandler loop device was employed to generate clots under shear (253 s−1) for testing, as previously described [46,47,48]. A rotating drum (110 mm diameter) was placed in a 37 °C water bath and set to rotate at a constant speed of 20 RPM. Tubing was cut to fit the drum diameter with an approximately 5 mm overlap and connectors prepared. A total of 2 mL of blood per clot formed was brought to a concentration of 1.5 million particles/mL of either 2, 5, 7, or 10 µm nPS particles. The blood was then recalcified by adding CaCl2 solution to a final concentration of 11 mM CaCl2. The recalcified blood was briefly mixed and loaded into the tubing using a syringe and placed on the drum to rotate for 1 h. After the clotting was complete the serum and clot were decanted from the tubing and the clot and tubing were rinsed with 1 mL of phosphate-buffered saline (PBS) solution. These rinses were added to the serum sample to ensure maximal particle recovery. The clot was then blotted to remove excess liquid, weighed, and added to a separate glass vial (see Supplementary Protocol S2).

2.6. Tissue Digestion

Tissue digestion was achieved by adding 1 mL of 10% KOH solution to each clot sample and 500 µL to each serum sample. Sample vials were mixed and then incubated overnight in a 50 °C water bath. The following day, samples were brought to a final concentration of 50 mM potassium phosphate buffer and brought to a neutral pH (7–8) with 5N HCl. Then, 100 µg/mL DNase was added, and the samples were allowed to incubate at 37 °C overnight. The next day SDS was added to bring the samples to 0.5% SDS and 250 µg/mL Proteinase K for incubation at 50 °C overnight. After digestion was complete the clot samples were run through the MEF device as described in previous experiments, and the serum samples were run through the apparatus using a higher flow 1 µm filter pore size. Filters were washed and the analysis was performed as previously described, with the modification of an additional rinse step for the clot sample vials. After the digested clot was filtered, but before the filter was rinsed, 2 mL of warm 0.01% Tween 20 solution was added to the empty clot sample vial and sonicated for 3 min while rotating in a sonicating bath. This vial rinse sample was loaded onto the same membrane as the digested clot and the final membrane processed as previously described.

2.7. Statistical Analysis

All data represent means unless described in the text as representative results. All data were collected and processed using Microsoft Excel and statistical testing performed using R. Particle sizes, concentrations, and recoveries are presented as means ± standard deviations. Particle recovery percentages were calculated comparing the pre- and post-MEF-isolated particles for each of the particle types, sizes, and concentrations tested. Paired student t-tests were calculated and statistical significance was deemed to be a p-value  <  0.05, where appropriate.

3. Results

Fluorescent polystyrene particles, ranging in size from 100 nm to 10 µm, with either nonfunctionalized (nPS), carboxylated (cPS), or aminated (aPS) surface modifications, were isolated via the MEF apparatus equipped with a 50 nm polycarbonate membrane filter. Submicron particle sizes (100, 300, and 500 nm) were analyzed on the Nanosight LM10 NTA (Nanoparticle Tracking Analysis). This video-based particle tracking system leverages particle mobility quantification, allowing for the simultaneous measurement of individual particle size and particle solution concentration with an upper particle size limit of ~1 µm. Particles above 1 µm, referred to here as micron particles (2, 5, 7, and 10 µm), were analyzed for size by microscopy, and particle recovery concentrations were quantified using a hemacytometer. For ease of discussion, the manuscript is split into sections based on the submicron- and micron-sized particle analyses throughout.

3.1. Submicron Particle Analysis

3.1.1. Size Distribution

All purchased particle sizes were experimentally verified prior to use. All particle sizes demonstrated a size variability that was not consistent across either size or surface modifications. The expected size of 500 nm varied by an average of 469.3 ± 2.6 nm nPS, 464.0 ± 5.1 nm aPS, and 385.9 ± 1.6 nm cPS. The expected size of 300 nm for the particles was measured, with averages of 293.1 ± 2.8 nm nPS, 311.8 ± 9.8 nm aPS, and 254.7 ± 2.5 nm cPS. The expected size of 100 nm varied, with averages of 107.1 ± 0.6 nm nPS, 102.7 ± 1.0 nm aPS, and 126.3 ± 4.6 nm cPS. The cPS particles showed the greatest deviation from the expected measurements, with those of 100 nm showing the highest percent deviation, being 26.3% larger, followed by the 500 nm cPS particles, which were 22.8% smaller (see Figure 2).
Aggregation was observed with the aPS and cPS particles when diluted into 0.01% Tween 20. This was largely resolved for the aPS and larger cPS particles by protein coating of the particles with 2 mg/mL BSA. However, the protein coating did not fully resolve the aggregation issue for the 100 nm cPS particles, with some remaining aggregation being observed throughout the experiments.
Because of the force exerted on the particles by the MEF process, the size was checked for the same sample before and after filtration to ensure there was no size change caused by deformation of the particles (see Supplementary Figure S1). A paired student t-test on all submicron particle groupings before and after filtration confirmed no significant difference in particles compared pre- and post-MEF isolation (p = 0.84, df = 24).

3.1.2. MEF Particle Recovery Varying Particle Size

Submicron particles sized 100, 300, and 500 nm with nonfunctionalized, aminated, and carboxylated surface modifications were filtered using a 50 nm filter on the mini-extruder, recovered from the filter using 0.01% Tween 20, and measured on the Nanosight LM10. Each condition was prepared in triplicate and compared to the same dilution that was not filtered. The percent recovery was assessed by calculating the change in particle concentration comparing the mini-extruded sample to its corresponding control dilution. All particle types were diluted to approximately the same concentration of 1 × 109 particles/mL. This concentration was chosen as it is at the high end of the Nanosight reading range of 107 to 109 particles/mL, which allows for poor recovery samples to remain within the quantitative reading range for the instrument. The average percent recovery for the 500 nm was 65.3% ± 7.1 for nPS, 98.3% ± 6.2 for aPS, and 78.2% ± 3.8 for cPS. These larger particles showed the strongest recovery, with the recovery decreasing as the size decreased. The 300 nm particle recovery followed this trend with 77.1% ± 8.9 for nPS, 61.2% ± 10.2 for aPS, and 80.8% ± 7.8 for cPS. The 100 nm particles showed the worst recovery, with an average percent recovery of 27.6% ± 2.7 for nPS, 7.0% ± 2.74 for aPS, and 16.3% ± 3.7 for cPS (see Figure 3).
While size had a strong effect on recovery, the surface modification had no clear trend in increasing or decreasing recovery. The aminated particles had the most linear relationship between size and recovery (R2 of 0.98) and, in the absence of other differentiating factors, was chosen as the particle type for further concentration recovery experiments.

3.1.3. MEF Particle Recovery Varying Particle Concentration

To examine how mini-extruder recovery is affected by the total particle load on the filter, samples were prepared using aPS particles in concentrations between 6 × 107 and 2.2 × 109 particles/mL (Figure 4). This range was chosen to cover the entire manufacturer-recommended particles-per-frame reading range of the Nanosight LM10. It is important to note that less concentrated and more concentrated samples can be analyzed via the Nanosight LM10 by simply concentrating or diluting the samples prior to analysis. Size continued to have a strong relationship with the recovery efficiency, with larger 500 nm particles showing consistent recovery across the tested concentration range (4.6 × 108 to 2.1 × 109 particles/mL). The 300 nm particles had a lower recovery overall, as well as a greater spread in recovery, with no observable trend in recovery across the concentrations. The 100 nm particles were the least consistent, showing the lowest average recovery rate and the greatest spread among the samples (see Figure 4).

3.2. Micron Particle Analysis

3.2.1. Size Distribution

Particles larger than one micron were sized using microscopy on a Nikon Eclipse Ni upright microscope (Melville, NY, USA) using the 20× objective for 5, 7, and 10 µm particles and 40× objective for 2 µm particles. Image analysis was carried out using QuPath software v0.5.0. According to this analysis, the majority of individual particle types and sizes were within 15% of the expected measurement, with some outliers being observed for the nPS and aPS particles, which were 23% and 28% larger than expected, respectively (see Figure 5, Supplementary Figure S2). Aggregation occurred when particles were allowed to settle for a significant period prior to loading into the hemacytometer; this aggregation was eliminated when particles were sonicated in a bath sonicator prior to loading.

3.2.2. MEF Particle Recovery Varying Particle Size

The micron-sized particle range was handled similarly to the submicron particles with stocks diluted into 2 mg/mL BSA and 0.01% Tween 20 in triplicate and loaded onto the MEF device at a set initial concentration between 105 and 106 particles/mL for the most accurate counts based on the recommended concentration range for the hemacytometer. Each sample replicate was read twice, and the average value for each sample was used to calculate the particle concentration of the recovered samples. The percent recovery was assessed by quantifying the change in particle concentration, comparing the MEF-isolated sample to each samples’ respective control dilution not run through the MEF device. For all particle surface modifications and particle sizes in the micron range, the recovery was high, with the highest recovery being 93.1 ± 0.8% observed for the 10 µm nPS particles and the lowest at 79.1% ± 9.9 for the 10 µm cPS particles. At the tested concentration of ~4.5 × 106 particles/mL, all of the micron particles across all of the surface modifications performed similarly, achieving an overall average recovery of 86.8% ± 4.3 across all conditions (see Figure 6A).

3.2.3. MEF Particle Recovery Varying Particle Concentration

Concentration-based effects on recovery were tested using the aPS particles, as there were no differentiating characteristics noted in the initial micron-sized particle recovery experiments, and this matched the submicron particle concentration experiment detailed previously. The range of tested concentrations varied among sizes, as each sample was compared directly to a concurrently prepared control dilution. The recommended quantitative range for the hemacytometer is 105 to 106 particles/mL, which was subsequently used as the range of concentrations tested for the micron-sized particle recovery experiments. Similar to the submicron 500 nm particles, the effect of concentration on recovery and surface modification was minimal, with the MEF system exhibiting effective recovery across the tested range (86.8 ± 4.3%, see Figure 6B). Environmental and biological plastic particle contaminants do not commonly occur as a single particle size, rather they are present in mixed-size groupings. To test the efficacy of the MEF particle recovery on the mixed particle samples, 5, 7, and 10 µm particles were mixed at approximately 3 × 106 particles/mL each. The mixed particle sample was then filtered and analyzed as previously described relative to a prefiltration control. The mixed samples achieved an average recovery of ≥90.7 ± 8.7 (see Supplementary Figure S3).
The entire suite of particles tested shows that isolation with the MEF device can achieve approximately 80% recovery of particles in a BSA and detergent solution from a size range of 500 nm to 10 µm and a greater than 60% recovery of particles down to 300 nm, with concentrations of 105 to 106 particles/mL for the micron-sized particle range and 107 to 109 particles/mL for submicron particles. (see Table 1).

3.3. Recovery from Digested Tissue

Up to this point, all particle recovery testing described herein using the MEF device occurred on purified pristine particles in liquid solutions, mimicking microplastic particle isolation and recovery from sources such as sea water or extracted environmental samples. The particle recovery efficiency from more complex digested biological tissues is also of critical importance and was, therefore, tested using the MEF system. As emerging research has identified microplastics in a variety of human tissue samples, a microplastic-infused human blood clot model was elected as a representative model system. Micron-sized (2, 5, 7, and 10 µm) nPS particles at a final concentration of 1.5 × 106 particles/mL were spiked into human blood and allowed to clot under shear flow using a Chandler loop apparatus in a 5/36″ ID 7/36″ OD tubing while rotating at 20 rpm for 1 h [46,47,48]. Following clot formation the formed clots, ranging in mass from 45.0 to 60.1 mg, were separated from the serum. There was no trend observed in mass related to particle size (average clot mass of 2 µm = 55.3 ± 4.5 mg, 5 µm = 49.0 ± 3.9 mg, 7 µm = 54.6 ± 5.3 mg, and 10 µm = 48.9 ± 3.0 mg). Potassium hydroxide (KOH) is capable of liquifying tissue while not damaging polystyrene [49] and was, therefore, selected as the initial digestion step for both the clots and serum followed by a DNase and Proteinase K enzymatic digestion to ensure complete fragmentation of large biological particles (see Supplementary Figure S4). Particle stability to elevated temperatures and concentrated KOH should be verified when utilizing other plastic particle types. The resulting digested liquid samples were neutralized with HCl and then filtered on the MEF apparatus using 0.4 µm polycarbonate filters. The percent recovery was assessed by comparing the final measured concentration to the calculated loaded particle concentration (see Figure 7).
The 5, 7, and 10 µm particles showed recoveries of ≥78 ± 11.4% consistent with the recovery results of the BSA/Tween buffer tests performed with the pristine micron particles. The recovery of 10 µm particles was highest with samples hitting 91 ± 9.4% recovery, followed by 7 µm at 87 ± 8.1% and 5 µm at 78 ± 11.4%. The 2 µm particles did not follow the trend and exhibited a recovery of only 9 ± 7.9%. The recovery of particles from the serum fractions were also assessed, with no particles being observed from the 5, 7, and 10 µm serum samples. However, there were particles recovered from the 2 µm serum fraction, exhibiting a recovery of 14.4 ± 14% of the loaded particles. This high particle recovery variability and high percentage of unaccounted for particles in the 2 µm clot and serum samples implies significant loss of particles to the Chandler loop tubing during the clot formation process. These results differed from the results observed in the 2 µm particle recovery from the less complex PBS and BSA buffer solutions.

4. Discussion

The purpose of the development of this nano-/microplastic particle-capture technique is to fill a gap in the recovery of particles between 100 nm and 10 µm in diameter that is currently unmet in the field. The MEF device provides for a simple and cost-effective particle filtration device that is both accessible and efficient. Different polystyrene particle sizes (100 nm to 10 µm) and surface chemistries (unmodified, carboxylated, and aminated) were tested, with the surface chemistries having less impact on particle recovery compared to the particle size.
Neither the submicron- nor the micron-sized particle ranges tested showed any recovery difference comparing the aminated, carboxylated, or unmodified surfaces. The lack of effects associated with surface modifications, which has been observed in the literature to be important to the reactivity and behavior of particles in vivo [50,51,52], is likely due to the BSA protein and Tween 20 coating incubation step used throughout this manuscript to prevent the aggregation of particles. The BSA exposure to the particles in the dilute sample fractions helped modulate the surface charge effects to both enhance particle recovery from the filter surface and reduce particle aggregation in the solution. Pristine, uncoated particles are unlikely to be found in environmental or biological samples, as protein coating is commonly observed due to ionic interactions between biological molecules and the plastic particle surfaces. For this reason, the use of BSA coating in the pristine polystyrene particle recovery tests described here provides for a representative purification of particles that may be found in environmental or sea water samples. For biologically derived particle samples, such as the microplastic-spiked blood clots described here, the addition of BSA in the liquid particle solution was not necessarily due to the presence of significant protein levels found within the sample tissue itself during the digestion process.
The size ranges tested were split into submicron (nanoplastics) and micron (microplastics) due to process-related differences, as they required different dilutions, and the particle recoveries were measured using different techniques. The micron-sized particles, 2 to 10 µm, exhibited strong particle recovery with an overall average of 86.8 ± 4.3% across all sizes and modifications at the same concentration and 84.1 ± 9.7% for all micron-sized aPS particles tested across the concentration range. These results demonstrate the consistent recovery of a variety of micron-sized particles at concentrations of 105 to 106 particles/mL exhibiting highly diverse surface charges.
The 500 nm particles similarly demonstrated a strong recovery efficiency; however, the 300 and 100 nm submicron particles’ recovery rate suffered with a decrease in particle sizes. The same aggregate average recovery across all submicron particles was 62.3 ± 30.6% for all sizes at a single concentration and 54.2 ± 33.8% across the concentration range of 107 to 109 particles/mL. Good recovery from the 300 nm particles was variable, with some samples achieving recovery rates similar to the 500 nm particles; however, the recovery consistency was lower than the larger particles. The 100 nm particles showed both lower recovery than the other samples and were highly inconsistent. For this reason, the current filter and protocol would not be recommended for 100 nm particles. The reduced recovery rate for the 100 nm particles was primarily driven by an inability to efficiently remove the successfully captured particles from the membrane itself following filtration. The smaller particles exhibited significantly higher membrane sticking than larger particles. The recovery inconsistency at smaller submicron particle sizes poses a potentially significant issue when applying this MEF technique to unknown plastic particle sample concentrations. It is likely that through continued optimization the recovery of submicron particles using the MEF system could be improved.
Recovery of 500 nm to 10 µm particles is effective from 105 through 106 particles/mL for particles in the micron range and between 107 and 109 particles/mL for the submicron particles. This testing range was limited by the instruments used to measure the particle counts. It is important to note that less concentrated and more concentrated samples can be analyzed via the Nanosight LM10 and hemacytometer by simply concentrating or diluting the sample prior to analysis. While not explicitly tested in this manuscript, the MEF device itself could also be used as an efficient particle concentrating platform by simply applying multiple 1 mL syringe volumes through the same membrane filter followed by recovery of the particles from the filter using a volume less than the original total sample volume.
Despite the utility of the mini-extruder filtration system as a quantitative capture method that is nondestructive and allows for multiple downstream analysis, there remain some limitations. Measurement and assessment of very small particles remain a challenge even after particles are effectively isolated. The sensitivity of the particle sizing and concentration measurement is limited by the sensitivity of the available assessment tools. Measuring particles in the submicron and micron range required the use of two separate instruments, and environmental/biological microplastics can occur in sizes ranging from several hundreds of microns to 50 nm or less. These particularly heterogenous samples might require sequential filtering with a larger pore size membrane filter in the MEF device to isolate micron and submicron particles from the same sample in two steps, which may result in further loss of the smaller particles. Membrane fouling is also a concern with any filtration system, and the very small pore sizes of the membranes used with the MEF device for the capture of submicron particles make the measurement of more complex solutions, such as cell digest, difficult, as seen in the preliminary clot digestion results presented here. Some particle recovery loss was observed to have occurred in the movement of samples between vials and to the inner surfaces of the MEF device itself; however, the most significant sources of unrecoverable particles were particles that were unable to be released from the filter surface. Removal of plastic particles using hydrophobic adsorption onto a filter is a known capture mechanism, so some difficulty removing particles bound to a filter is expected depending upon the filter composition and the solution from which the particles are purified [44,45,53]. Sonication and the use of warm buffers assisted in particle release from the filter, in addition to avoiding particles becoming tightly attached due to high pressures when loading the sample onto the membrane. Different concentrations of detergent for washing the filter were tested, but higher concentrations (>0.01% Tween 20) provided minimal benefit and began to interfere with the fluorescent signaling of the submicron particles. Higher concentrations of BSA were not observed to cause significant improvement in recovery, and the much more proteinaceous digested blood clot solution performed similarly to 2 mg/mL BSA for all but the 2 µm particles. Finally, this system is best suited for smaller volumes. There is no upper limit to the volume that can be filtered, but practically anything larger than a few milliliters would be difficult to process and may require a preprocessing concentration step.

5. Conclusions

Use of the MEF platform for filtration-based isolation of plastic particles from solution allows for an accessible means of extending the lower limit of nano- and microplastic particles that can be recovered through a simple and reproducible isolation technique. Nanometer- and small micron-sized plastic particles show some of the strongest detrimental effects to living organisms, with their presence and characteristics in the environment and biological samples being the most difficult to isolate, quantify, and analyze. This study utilized spherical polystyrene (PS) particles as model nano- and microplastic MEF-isolation validation particle types; however, environmental and biological mico- and nanoplastics are commonly fragmented and irregular. Future work using the MEF device includes exploring the use of different filters, the use of particles with more complex surface characteristics, and assessing different polymer types and shapes which may change the dynamics of the MEF filter loading. Further testing is also needed on filtration of particles from complex samples, such as other digested animal and human tissue types.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/environments11080180/s1, Figure S1: Representative size distribution data for submicron particle before and after MEF isolation; Figure S2: Representative microscope sizing for aPS and cPS micron particles; Figure S3: Mixed 5, 7, and 10 µm particle recovery; Figure S4: Representative images of KOH clot digestion process; Protocol S1: MEF protocol; Protocol S2: Chandler loop clot formation protocol.

Author Contributions

Conceptualization: N.J.A.; Methodology: A.H., L.F.C.P., K.H. and A.C.; Validation: K.H. and A.C.; Formal analysis: A.H. and L.F.C.P.; Investigation: A.H. and L.F.C.P.; Resources: N.J.A.; Data curation: A.H. and L.F.C.P.; Writing—original draft preparation: A.H., L.F.C.P. and N.J.A.; Supervision: N.J.A.; Project administration: N.J.A. All authors have read and agreed to the published version of the manuscript.

Funding

This project was funded, in part, with support from the Indiana Clinical and Translational Sciences Institute Grant, [UL1TR002529 (IMPRS) from the National Institutes of Health] and, in part, by the Louis Stokes STEM Pathways Implementation-Only Alliance: Indiana LSAMP [NSF EES 2308500 (SPIO Indiana LSAMP)] from the National Science Foundation. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health or National Science Foundation.

Data Availability Statement

The raw data supporting the conclusions of this article will be made available by the authors upon request.

Acknowledgments

Figures were made with BioRender.com.

Conflicts of Interest

The authors declare no conflicts of interest.

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Figure 1. (A) Schematic overview of the experimental steps, including capture of particles onto polycarbonate filters in the mini-extruder, recovery by rinsing and sonication in glass vials, and then analysis with either the Nanosight LM10 for submicron-sized particles or hemacytometer for micron-sized particles. (B) Labeled picture of the mini-extruder filtration (MEF) device and filter orientation.
Figure 1. (A) Schematic overview of the experimental steps, including capture of particles onto polycarbonate filters in the mini-extruder, recovery by rinsing and sonication in glass vials, and then analysis with either the Nanosight LM10 for submicron-sized particles or hemacytometer for micron-sized particles. (B) Labeled picture of the mini-extruder filtration (MEF) device and filter orientation.
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Figure 2. (A) A representative frame of video captured by the Nanosight LM10 of 500 nm nPS particles. Representative Nanosight sizing distributions for submicron (B) nPS, (C) aPS, and (D) cPS particles. All particle sizing data are an average of three measurements of a single sample. Concentrations across particle sizes are not normalized.
Figure 2. (A) A representative frame of video captured by the Nanosight LM10 of 500 nm nPS particles. Representative Nanosight sizing distributions for submicron (B) nPS, (C) aPS, and (D) cPS particles. All particle sizing data are an average of three measurements of a single sample. Concentrations across particle sizes are not normalized.
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Figure 3. Percent recovery assessed at similar concentrations for the aPS, cPS, and nPS particles measured at 0.1, 0.3, and 0.5 µm in diameter.
Figure 3. Percent recovery assessed at similar concentrations for the aPS, cPS, and nPS particles measured at 0.1, 0.3, and 0.5 µm in diameter.
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Figure 4. Percent recovery for submicron aPS particles across different concentrations.
Figure 4. Percent recovery for submicron aPS particles across different concentrations.
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Figure 5. Representative brightfield microscope images of nPS (A) 2, (B) 5, (C) 7, and (D) 10 µm particles including average ± standard deviation sizing data. All particles included a minimum of 50 individual particle measurements.
Figure 5. Representative brightfield microscope images of nPS (A) 2, (B) 5, (C) 7, and (D) 10 µm particles including average ± standard deviation sizing data. All particles included a minimum of 50 individual particle measurements.
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Figure 6. (A) Percent recovery assessed at similar concentrations for aPS, cPS and nPS particles measured between 2 and 10 µm in diameter. (B) Particle recovery of aPS micron-sized samples isolated on the MEF device across a range of particle concentrations.
Figure 6. (A) Percent recovery assessed at similar concentrations for aPS, cPS and nPS particles measured between 2 and 10 µm in diameter. (B) Particle recovery of aPS micron-sized samples isolated on the MEF device across a range of particle concentrations.
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Figure 7. (A) Representative clot before digestions; (B) clot after incubation in 10% KOH at 50 °C for 24 h; (C) percent recovery of micron-sized particles from digested human blood clots after filtration.
Figure 7. (A) Representative clot before digestions; (B) clot after incubation in 10% KOH at 50 °C for 24 h; (C) percent recovery of micron-sized particles from digested human blood clots after filtration.
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Table 1. All recovery results for all particle sizes and surface modifications tested. A loading volume of 400 µL on the MEF device was used for all samples.
Table 1. All recovery results for all particle sizes and surface modifications tested. A loading volume of 400 µL on the MEF device was used for all samples.
Particle TypeSize (µm)Particles × 106/mLPercent Recovery
aPS0.113307.0 ± 2.7
0.3107061.2 ± 10.2
0.5168098.3 ± 6.2
2.04.491.1 ± 12.4
5.05.788.5 ± 3.2
7.04.691.4 ± 6.2
10.03.386.6 ± 6.2
cPS0.1132016.3 ± 3.7
0.3198080.8 ± 7.8
0.5160078.2 ± 3.8
2.04.690.8 ± 6.2
5.04.384.7 ± 7.7
7.04.084.3 ± 5.6
10.04.379.1 ± 9.9
nPS0.1206027.6 ± 7.4
0.3207077.1 ± 8.9
0.5175065.3 ± 7.1
2.04.384.0 ± 9.3
5.04.582.1 ± 4.5
7.04.685.9 ± 12.3
10.04.993.1 ± 0.8
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MDPI and ACS Style

Hall, A.; Cardona Polo, L.F.; Helms, K.; Christodoulides, A.; Alves, N.J. Microplastic and Nanoplastic Particle Isolation from Liquid and Biological Samples via Mini-Extruder Filtration (MEF). Environments 2024, 11, 180. https://doi.org/10.3390/environments11080180

AMA Style

Hall A, Cardona Polo LF, Helms K, Christodoulides A, Alves NJ. Microplastic and Nanoplastic Particle Isolation from Liquid and Biological Samples via Mini-Extruder Filtration (MEF). Environments. 2024; 11(8):180. https://doi.org/10.3390/environments11080180

Chicago/Turabian Style

Hall, Abigail, Luis F. Cardona Polo, Kennedy Helms, Alexei Christodoulides, and Nathan J. Alves. 2024. "Microplastic and Nanoplastic Particle Isolation from Liquid and Biological Samples via Mini-Extruder Filtration (MEF)" Environments 11, no. 8: 180. https://doi.org/10.3390/environments11080180

APA Style

Hall, A., Cardona Polo, L. F., Helms, K., Christodoulides, A., & Alves, N. J. (2024). Microplastic and Nanoplastic Particle Isolation from Liquid and Biological Samples via Mini-Extruder Filtration (MEF). Environments, 11(8), 180. https://doi.org/10.3390/environments11080180

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