Submitted Successfully!
To reward your contribution, here is a gift for you: A free trial for our video production service.
Thank you for your contribution! You can also upload a video entry or images related to this topic.
Version Summary Created by Modification Content Size Created at Operation
1
Andrea Masseroni
 -- 2447 2022-06-13 04:27:58 |
2 Format correction
Sirius Huang
 Meta information modification 2447 2022-06-13 04:33:16 |

Video Upload Options

Do you have a full video?
Send video materials Upload full video

Confirm

Are you sure to Delete?
Yes No
Cite
If you have any further questions, please contact Encyclopedia Editorial Office.
Masseroni, A.; Villa, S.; Rizzi, C.; Urani, C. Exposure Assessment of Nanoplastics. Encyclopedia. Available online: https://encyclopedia.pub/entry/23950 (accessed on 27 July 2024).
Masseroni A, Villa S, Rizzi C, Urani C. Exposure Assessment of Nanoplastics. Encyclopedia. Available at: https://encyclopedia.pub/entry/23950. Accessed July 27, 2024.
Masseroni, Andrea, Sara Villa, Cristiana Rizzi, Chiara Urani. "Exposure Assessment of Nanoplastics" Encyclopedia, https://encyclopedia.pub/entry/23950 (accessed July 27, 2024).
Masseroni, A., Villa, S., Rizzi, C., & Urani, C. (2022, June 13). Exposure Assessment of Nanoplastics. In Encyclopedia. https://encyclopedia.pub/entry/23950
Masseroni, Andrea, et al. "Exposure Assessment of Nanoplastics." Encyclopedia. Web. 13 June, 2022.
Exposure Assessment of Nanoplastics
Edit
This entry is adapted from the peer-reviewed paper 10.3390/toxics10050270

Nanoplastics (NPs) are particles ranging in size between 1 and 1000 nm, and they are a form of environmental contaminant of great ecotoxicological concern. The detection of these contaminants in complex matrices is a real challenge. Developing suitable and reliable analytical methods for quantifying the environmental occurrences of NPs is pivotal. 

nanoplastics environmental risk assessment effects exposure

1. Introduction

Very limited scientific literature regarding the occurrence and concentration of Nanoplastics (NPs) in the environment is available. The detection of these contaminants in complex matrices is a real challenge. As such, the nanometric size of NPs renders the common extractive approaches applied for MPs unsuitable [1], and the heterogeneity of NP polymers makes the application of techniques that are typically used for engineered nanomaterials (ENMs) difficult [2]. Contrary to ENMs, which present a uniform composition owing to their intentional production as nanoparticles, NPs contain a mixture of different polymer types, sizes, and shapes, because they are derived from the unintentional fragmentation of larger plastic items dispersed in the environment. Moreover, the presence of a large variety of additives in the plastic materials of origin hinders the identification of polymer [1][2][3].
Currently, there are no straightforward methodologies for the extraction, identification, and quantification of NPs in the environment [4]. Eight studies have successfully extracted these contaminants from abiotic field samples. These studies have documented the occurrence of nanoscale plastic particles in seawater [5], rivers [6], snow [7][8], air [9], soil [10], sand [11], and tap water [12]. As shown in Table 1, only a few studies have simultaneously reported the concentration and relative polymeric types of NPs present in samples [6][7][12].
Table 1. List of the studies that successfully detected NPs in environmental abiotic matrices (updated to December 2021). PA: polyamide; PO: polyolefins.
Samples Concentration Polymer Type References
Seawater - PET, PS, PE, PVC [5]
River water 1.92–2.82 μg/L PS [6]
Snow 46.5 μg/L PET, PP [7][8]
Air - PET, PS [9]
Soil - PVC, PS, PE [10]
Sand - PVC, PS [11]
Tap water 1.67–2.08 μg/L PVC, PS, PA, PO [12]
The measured environmental concentrations of NPs are extremely low (μg·L−1), posing significant analytical challenges for their detection in the environment. A possible strategy to overcome some of these limitations is to consider the use of biomonitoring. Bioindicators are living organisms that provide information on environmental quality [13]. More precisely, exposure indicators (e.g., bioaccumulation of xenobiotics in tissues) are considered useful tools to obtain data on environmental pollutants [14].
Owing to their capacity to accumulate nanosized particles, organisms may accumulate higher concentrations of NPs in the biological matrix than in other complex matrices [15]. Following uptake, NPs, especially the small ones (<100 nm), can penetrate the biological membranes [16] and potentially bioaccumulate in tissues or organs, leading to their transfer along trophic chains. This assumption is supported by experimental evidence [17]. Briefly, Zhou et al. [17] reported for the first time that the concentration of NPs in aquatic organisms is approximately 1 μg·g−1. Therefore, the use of biomonitoring offers significant advantages. Specifically, once the most suitable biomonitor is selected for different matrices, the extraction methodologies for different organisms only require optimization at the digestion step. As with any novel approach to a challenging issue, biomonitoring presents critical points. Assuming that NP concentration can be measured in biomonitor tissues, composition of the medium to which organisms are exposed must be estimated. For classical contaminants, the conventional approach is to achieve medium exposure through partition coefficients (e.g., Kow). However, the factors driving NP bioaccumulation and/or bioconcentration remain largely unknown. Therefore, the correct approach to calculating NP concentrations in the environmental matrix in which the organisms live remains unknown. Notwithstanding, a promising approach to model the mass balance of ingestion and loss processes of MPs has been recently reviewed [18], highlighting the possibility of modelling NP uptake and release in a similar manner. To calibrate and validate models, experimental determination of concentration is pivotal; consequently, a sound methodology is essential.
Eight studies to date have proposed different methodologies to achieve this goal [15][17][19][20][21][22][23][24] (Table 2). All these studies utilized non-properly ‘environmental’ organisms, since all samples were collected from local markets [15][17][19] or directly from farms [20][21]. To test the efficiency of the proposed methodologies, tissue samples were spiked with nanoPS. Zhou et al. [17] verified the suitability of their method in Tilapia sp. and, as mentioned previously, successfully proved the environmental occurrence of NPs in three different species, at concentrations ranging from 0.09 to 0.78 μg·g−1.
Table 2. List of studies that developed novel methodologies for the detection of NPs in biological samples.
Organism Target Polymer Size (nm) Ref.
Mollusk, Crustacean, Fish Muscle PS, PMMA 100 [17]
Mollusk (oyster)
Mammal (C57BL/6 mice)		 Whole organism
Gut, liver, kidney		 PS 70
70		 [19]
Fish (D. labrax) Muscle PS 100 [20]
Bird (non-specified) Eggshell PS 60, 200, 600 [21]
Mammal (R. norvegicus) Blood PS 100, 200, 500 [22]
Tunicate (R. ciona) Whole organism PS 100 [15]
Mollusk (M. galloprovincialis) Whole organism PS 100, 500, 1000 [23]
Mollusk (M. edulis) Stomach PS 49 [24]
The application of such methodologies can aid better comprehension of NP exposure in the environment. Nonetheless, the detection of NPs in biological samples is limited by certain analytical issues. From this perspective, a harmonized methodology of biological extraction is required to favor intercomparisons amongst studies. In the following sections, innovations reported in studies listed in Table 3 are elaborated, comparing different proposed extraction methods, and highlighting their strengths and critical points.

2. Digestion of Organic Matter in Biological Samples

Steps required for the extraction of NPs from biological samples can be summarized as follows: sampling, pre-treatment, digestion, preconcentration, separation, identification, and quantification [4][17].
Regarding sampling methods, please refer to a recently published review on this topic [25]. The complexity of the biological matrix makes the detection of NPs challenging. Plastic particles must be isolated and separated from the organic matter of biological samples, which could lead to interference in the subsequent steps of extraction. In this perspective, digestion is a key step which should be suitable to remove the organic matter, preserving, however, the properties and quantity of NPs [1][26]. Two different approaches are commonly adopted: chemical degradation and enzymatic digestion [4][27]. The first approach involves the use of different chemicals for acid (e.g., HCl and HNO3) or alkaline (e.g., NaOH and KOH) digestion, whilst the second utilizes specific enzymes (e.g., proteinase K, papain) that degrade the organic matrix. Chemical degradation enables efficient removal of the background matrix and is a cost-effective approach. However, acid treatment has recently been questioned because it tends to destroy and cause aggregation of NPs polymers [20][27]. Therefore, an alkaline approach, which is less invasive, is recommended [22]. Zhou et al. [17] obtained satisfactory results using this approach to digest the organic tissues of fish samples. When comparing the efficiency of two different alkaline reagents, tetramethylammonium hydroxide (TMAH) and sodium hydroxide (NaOH), TMAH showed a greater recovery without NP aggregation. In another recent study, Muhammad et al. [28] digested the gut and intestinal contents of insect larvae following the digestion protocol proposed by Zhou et al. [19].
Meanwhile, the enzymatic approach is considered a valid alternative to alkaline degradation. Although this approach incurs higher costs [25], it is non-destructive and avoids particle aggregation [29]. Correia and Loeschner [20] compared acid degradation (HNO3) with enzymatic digestion (proteinase K) in muscle tissues of fish samples and demonstrated that the former approach led to NP aggregation and subsequent hindrance in the extractive steps, whilst the latter could overcome these analytical issues. Furthermore, a recently proposed enzymatic digestion protocol [29] has shown satisfactory results for NPs extraction from biological matrices [15][23].

3. Separation, Identification, and Quantification of NPs in Biological Samples

Following digestion, it is necessary to isolate and separate particles in the nanometric size range; identify their effective sizes, shapes, and concentrations, and confirm the effective plastic nature of the extracted polymers through chemical characterisation. Simultaneous achievement of these results remains a huge challenge because of the lack of an efficient methodology for the extraction, identification, and quantification of NPs. The main steps adopted in the proposed methodologies for the detection of NPs in biological samples are summarized in Table 3.
Table 3. Methods for the detection of NPs in biological samples. TEM: transmission electron microscopy; SEM: scanning electron microscopy; py–GC–MS: pyrolysis–gas chromatography–mass spectrometry; CSE, coagulation-sedimentation extraction; AF4, asymmetrical flow-field fractionation; DAD, diode array detector; MALS, multiangle light scattering detector; CRM, confocal Raman spectroscopy; EDX, energy-dispersive X-ray; FIB, focused ion beam.
Digestion Separation Quantification Identification Ref.
Alkali (TMAH) Ethanol-precipitation TEM py-GC-MS [17]
Alkali (KOH) CSE - py-GC-MS [19]
Enzimatic (Proteinase K) AF4 AF4-MALS - [20]
Acid (HCl)
Alkali (TMAH)		 AF4
Ultracentrifugation		 AF4-MALS - [21]
Alkali
(KOH)		 AF4 AF4-DAD-MALS
TEM		 - [22]
Enzimatic (Papain) AF4, Ultrafiltration
Chip trapping		 AF4-DAD-MALS
SEM		 CRM
SEM-EDX		 [15]
Enzimatic (Papain)
FIB		 Microcavity-size selection SEM CRM
SEM-EDX		 [23]
Alkali (NaOH, KOH) Centrifugation Microplate fluorescence reader - [24]

3.1. NP Precipitation

Two different methodologies have been proposed to separate and isolate NPs present in biological samples [17][19] by exploiting the tendency of NPs to aggregate and co-precipitate with suspended materials. Typically, NPs tend to be dispersed in the medium, since they are dominated by Brownian motion; however, under specific conditions of pH and organic matter concentration, they associate with the material present in the solution, with their consequent sedimentation [19][30].
Zhou et al. [17] utilized the high binding affinity of NPs to proteins to extract them from biological samples through ethanol precipitation. The authors successfully validated the effectiveness of the proposed method by spiking the digested muscle tissue samples from Tilapia sp. with 2 μg·g−1 of PS and PMMA of different sizes (50, 100, and 500 nm) and obtained high recovery rates for all polymer sizes. Moreover, using pyrolysis–gas chromatography–mass spectrometry (py–GC–MS), they reported excellent limits of detections (LODs) (0.03 and 0.09 μg·g−1 for 100 nm PS and PMMA, respectively). Further, by adopting this methodology for environmental samples, the authors succeeded in detecting nanoPS (0.093–0.785 μg·g−1) in three different aquatic species.
Gao et al. [19] proposed a protocol based on coagulation–sedimentation extraction (CSE) between NPs and diatomite. By spiking the tissue samples from oysters and mice with nanoPS (70 nm), microPS (2 μm), and diatomite (7 μm), the authors successfully isolated and extracted NPs. As opposed to MPs, NPs tend to bind diatomite and precipitate upon centrifugation; thus, NPs can be isolated from MPs and the remaining suspended material with high recovery rates (95%). Finally, the authors chemically characterized the extracted particles using py–GC–MS and obtained an LOD of 0.012 μg·g−1.

3.2. AF4 and Chip Trapping

Various studies have adopted an approach based on asymmetrical flow-field fractionation (AF4) to separate NPs from spiked biological samples [20][21][22][23]. AF4 allows the fractionation of nanoparticles based on their hydrodynamic size, which upon combination with detectors, such as diode array detector (DAD) and multiangle light scattering detector (MALS), provides additional information on particle size and concentration [4][31].
The first studies that adopted this approach in biological samples performed AF4-MALS to separate NPs from spiked digested fish tissue [20] and bird shell samples [21]. Luo et al. [22] recently proposed an AF4-DAD-MALS approach to separate and detect nanoPS (from 30 to 500 nm) in spiked blood samples of rats (Rattus norvegicus), demonstrating the feasibility of this methodology to separate and quantify nanoPS in biological samples, with satisfactory results for >100 nm PS.
In a recent study, Valsesia et al. [15] proposed a methodology for separating biological sample fractions with the same particle size. The authors spiked tunicate samples (Ciona robusta) with nanoPS (100 nm) and subjected them to AF4-DAD-MALS following enzymatic digestion. Subsequently, through ultrafiltration, the AF4-derived fractions were concentrated to a small volume and placed on a chip. Exploiting the tendency of nanoparticles to aggregate as the sample dries, NP clusters were grouped in a small area of the chip, achieving a sufficient signal for analysis using confocal Raman spectroscopy (CRM). By adopting this procedure, the authors succeeded in the chemical characterisation of nanoPS, combined with the estimation of NP concentration using scanning electron microscopy (SEM) coupled with an energy-dispersive X-ray (EDX) analyzer. The proposed method yields concentration data that are comparable to environmental levels.
In another study, Valsesia et al. [23] proposed a similar methodology that involves drying a sample on a chip with arrays of cavities of different sizes. Owing to the peculiar structure of the chip surface, single nanoparticles fall into the cavities according to their size, allowing the formation of small NP aggregates. The precise positioning of NPs of a certain size in the corresponding nanocavities enables their examination using SEM and spectroscopy (CRM). By testing the methodology on mollusk tissues (Mytilus galloprovincialis) exposed to 100 nm nanoPS, the authors succeeded in isolating nanoPS using a chip with 300 nm pores on the surface and finally characterized the isolated particles using CRM.
Moraz and Breider [24], improving the previously proposed methods [32][33], proposed a fluorescence-based approach for the detection of nanoPS in biological tissues. The method uses a fluorescent probe (a particular molecular rotor) that can conjugate with NPs. However, owing to the complexity of the investigated biological matrix (Mytilus edulis tissues), the authors did not obtain satisfactory results; nonetheless, the methodology presents the feasibility of detecting and quantifying NPs in biological samples.

4. Comparison Amongst Different Approaches

By comparing the methodologies presented in the literature, attempts can be made to fill this gap for successful extraction, identification, and quantification of NPs in biological samples.
The extraction of NPs from biological samples requires an efficient digestion step that allows for the removal of organic matter without damaging or causing the agglomeration of NPs. From this perspective, enzymes (e.g., papain) and alkaline reagents (e.g., TMAH) appear to be the most promising. Once digestion is performed, several methodologies can be adopted. Currently, however, no straightforward approach that allows simultaneous collection of data on concentrations and polymer characterisation is available.
The precipitation approach, followed by py–GC–MS, is relatively simple, yields an excellent LOD (e.g., 0.03 μg·g−1 [17]) and a high recovery rate (>85% [17][19]), and allows for the detection of small NPs (e.g., nanoPS 70 nm, [19]). However, although the application of py–GC–MS allows for covering the entire nanometric range, it is a destructive method that makes it impossible to simultaneously obtain information on the physical properties of individual particles [4]. Furthermore, because this methodology involves size-selective precipitation, handling real samples containing NPs of different sizes may be difficult. From this perspective, the use of AF4 as a separation step may be an effective alternative for obtaining fractions with NPs of the same size.
Furthermore, methodologies based on the evaporation of sample droplets on specific chip surfaces, followed by CRM analysis, are non-destructive. Using these methods, the number of particles can be directly counted through SEM and their polymeric type can be directly confirmed through Raman spectroscopy, with satisfactory LODs. However, these approaches require advanced instrumentation (e.g., functionalized chips) and additional analytical steps, such as an added purification step involving focused ion beam (FIB) [23], ultrafiltration [15], and EDX analysis, which may result in greater reproducibility.
Owing to the complexity of NP pollution, formulation of an extraction protocol for these contaminants from biological samples warrants effort. However, the scientific community is moving in the right direction, with significant progress being made towards better understanding the effective concentrations and levels of NPs in the environment.

References

  1. Jakubowicz, I.; Enebro, J.; Yarahmadi, N. Challenges in the search for nanoplastics in the environment. A critical review from the polymer science perspective. Polym. Test. 2021, 93, 106953.
  2. Gigault, J.; El Hadri, H.; Nguyen, B.; Grassl, B.; Rowenczyk, L.; Tufenkji, N.; Feng, S.; Wiesner, M. Nanoplastics are neither microplastics nor engineered nanoparticles. Nat. Nanotechnol. 2021, 16, 501–507.
  3. Nava, V.; Frezzotti, M.L.; Leoni, B. Raman Spectroscopy for the Analysis of Microplastics in Aquatic Systems. Appl. Spectrosc. 2021, 75, 1341–1357.
  4. Cai, H.; Xu, E.G.; Du, F.; Li, F.; Liu, J.; Shi, H. Analysis of environmental nanoplastics: Progress and challenges. Chem. Eng. J. 2021, 410, 128208.
  5. Ter Halle, A.; Jeanneau, L.; Martignac, M.; Jardé, E.; Pedrono, B.; Brach, L.; Gigault, J. Nanoplastic in the North Atlantic Subtropical Gyre. Environ. Sci. Technol. 2017, 51, 13689–13697.
  6. Zhou, X.X.; He, S.; Gao, Y.; Li, Z.C.; Chi, H.Y.; Li, C.J.; Wang, D.J.; Yan, B. Protein Corona-Mediated Extraction for Quantitative Analysis of Nanoplastics in Environmental Waters by Pyrolysis Gas Chromatography/Mass Spectrometry. Anal. Chem. 2021, 93, 6698–6705.
  7. Materic, D.; Kasper-Giebl, A.; Kau, D.; Anten, M.; Greilinger, M.; Ludewig, E.; van Sebille, E.; Rockmann, T.; Holzinger, R. Micro- and nanoplastics in alpine snow: A new method for chemical identification and (semi)quantification in the nanogram range. Environ. Sci. Technol. 2020, 54, 2353–2359.
  8. Materic, D.; Ludewig, E.; Brunner, D.; Rockmann, T.; Holzinger, R. Nanoplastics transport to the remote, high-altitude Alps (2021). Environ. Pollut. 2021, 288, 117697.
  9. Xu, G.; Cheng, H.; Jones, R.; Feng, Y.; Gong, K.; Li, K.; Fang, X.; Tahir, M.; Valev, V.; Zhang, L. Surface-Enhanced Raman Spectroscopy Facilitates the Detection of Microplastics <1 μm in the Environment. Environ. Sci. Technol. 2020, 54, 15594–15603.
  10. Wahl, A.; Le Juge, C.; Davranche, M.; El Hadri, H.; Grassl, B.; Reynaud, S.; Gigault, J. Nanoplastic occurrence in a soil amended with plastic debris. Chemosphere 2021, 262, 127784.
  11. Davranche, M.; Lory, C.; Le Juge, C.; Blancho, F.; Dia, A.; Grassl, B.; El Hadri, H.; Pascal, P.Y.; Gigault, J. Nanoplastics on the coast exposed to the North Atlantic Gyre: Evidence and traceability. NanoImpact 2020, 20, 100262.
  12. Li, Y.; Wang, Z.; Guan, B. Separation and identification of nanoplastics in tap water. Environ. Res. 2022, 204, 112134.
  13. Burger, J. Bioindicators: Types, Development, and Use in Ecological Assessment and Research. Environ. Bioindic. 2016, 1, 22–39.
  14. Messer, J.J.; Linthurst, R.A.; Overton, W.S. An EPA program for monitoring ecological status and trends. Environ. Monit. Assess. 1991, 17, 67–78.
  15. Valsesia, A.; Parot, J.; Ponti, J.; Mehn, D.; Marino, R.; Melillo, D.; Muramoto, S.; Verkouteren, M.; AHackley, V.A.; Colpo, P. Detection, counting and characterization of nanoplastics in marine bioindicators: A proof of principle study. Microplastics Nanoplastics 2021, 1, 5.
  16. Azeem, I.; Adeel, M.; Ahmad, M.A.; Shakoor, N.; Jiangcuo, G.D.; Azeem, K.; Ishfaq, M.; Shakoor, A.; Ayaz, M.; Xu, M.; et al. Uptake and Accumulation of Nano/Microplastics in Plants: A Critical Review. Nanomaterials 2021, 11, 2935.
  17. Zhou, X.X.; He, S.; Gao, Y.; Chi, H.Y.; Wang, D.J.; Li, Z.C.; Yan, B. Quantitative Analysis of Polystyrene and Poly(methyl methacrylate) Nanoplastics in Tissues of Aquatic Animals. Environ. Sci. Technol. 2021, 55, 3032–3040.
  18. Besseling, E.; Redondo-Hasselerharm, P.; Foekema, E.M.; Koelmans, A.A. Quantifying ecological risks of aquatic micro- and Nanoplastic. Environ. Sci. Technol. 2019, 49, 32–80.
  19. Gao, H.; Lin, Y.; Wei, J.; Zhang, Y.; Pan, H.; Ren, M.; Li, J.; Huang, L.; Zhang, X.; Huang, Q.; et al. A novel extraction protocol of nano-polystyrene from biological samples. Sci. Total Environ. 2021, 790, 148085.
  20. Correia, M.; Loeschner, K. Detection of nanoplastics in food by asymmetric flow field-flow fractionation coupled to multi-angle light scattering: Possibilities, challenges and analytical limitations. Anal. Bioanal. Chem. 2018, 410, 5603–5615.
  21. Monikh, F.A.; Grundschober, N.; Romeijn, S.; Arenas-Lago, D.; Vijver, M.G.; Jiskoot, W.; Peijnenburg, W.J.G.M. Development of methods for extraction and analytical characterization of carbon-based nanomaterials (nanoplastics and carbon nanotubes) in biological and environmental matrices by asymmetrical flow field-flow fractionation. Environ. Pollut. 2019, 255, 113304.
  22. Luo, X.; Wu, Y.; Zhang, L.; Li, K.; Jia, T.; Chen, Y.; Zhou, L.; Huang, P. An effective solution to simultaneously analyze size, mass and number concentration of polydisperse nanoplastics in a biological matrix: Asymmetrical flow field fractionation coupled with a diode array detector and multiangle light scattering. RSC Adv. 2021, 11, 12902–12906.
  23. Valsesia, A.; Quarato, M.; Ponti, J.; Fumagalli, F.; Gilliand, D.; Colpo, P. Combining microcavity size selection with Raman microscopy for the characterization of Nanoplastics in complex matrices. Sci. Rep. 2021, 11, 362.
  24. Moraz, A.; Breider, F. Detection and Quantification of Nonlabeled Polystyrene Nanoparticles Using a Fluorescent Molecular Rotor. Anal. Chem. 2021, 93, 14976–14984.
  25. Lai, Y.; Dong, L.; Li, Q.; Li, P.; Liu, J. Sampling of micro- and nano-plastics in environmental matrixes. Trends Anal. Chem. 2021, 145, 116461.
  26. Li, P.; Li, Q.; Hao, Z.; Yu, S.; Liu, J. Analytical methods and environmental processes of nanoplastic. Res. J. Environ. Sci. 2020, 94, 88–99.
  27. Cerasa, M.; Teodori, S.; Pietrelli, L. Searching Nanoplastics: From Sampling to Sample Processing. Polymers 2021, 13, 3658.
  28. Muhammad, A.; Zhou, X.X.; He, J.; Zhang, N.; Shen, X.; Sun, C.; Yan, B.; Shao, Y. Toxic effects of acute exposure to polystyrene microplastics and nanoplastics on the model insect, silkworm Bombyx mori. Environ. Pollut. 2021, 285, 117255.
  29. Facchetti, S.V.; La Spina, R.; Fumagalli, F.; Riccardi, N.; Gilliland, D.; Ponti, J. Detection of Metal-Doped Fluorescent PVC Microplastics in Freshwater Mussels. Nanomaterials 2020, 10, 2363.
  30. Sun, H.; Jiao, R.; Wang, D. The difference of aggregation mechanism between microplastics and nanoplastics: Role of Brownian motion and structural layer force. Environ. Pollut. 2021, 268, 115942.
  31. Enyoh, C.E.; Wang, Q.; Chowdhury, T.; Wang, W.; Lu, S.; Xiao, K.; Chowdhury, M.A.H. New Analytical Approaches for Effective Quantification and Identification of Nanoplastics in Environmental Samples. Processes 2021, 9, 2086.
  32. Bisso, P.W.; Tai, M.; Katepalli, H.; Bertrand, N.; Blankschtein, D.; Langer, R. Molecular Rotors for Universal Quantitation of Nanoscale Hydrophobic Interfaces in Microplate Format. Nano Lett. 2018, 18, 618–628.
  33. Gagné, F. Detection of polystyrene nanoplastics in biological tissues with a fluorescent molecular rotor probe. J. Xenobiot. 2019, 9, 8147.
More
© Text is available under the terms and conditions of the Creative Commons Attribution (CC BY) license; additional terms may apply. By using this site, you agree to the Terms and Conditions and Privacy Policy.
Upload a video for this entry
Information
Subjects: Toxicology; Environmental Sciences
Contributors MDPI registered users' name will be linked to their SciProfiles pages. To register with us, please refer to https://encyclopedia.pub/register :
Andrea Masseroni
,
Sara Villa
,
Cristiana Rizzi
,
Chiara Urani
View Times: 396
Revisions: 2 times (View History)
Update Date: 13 Jun 2022
Table of Contents
    1000/1000
    Hot Most Recent
    Video Production Service
    Feedback