Contaminants of Emerging Concern on Microplastics Found in the Chrysaora chesapeakei of the Patuxent River, Chesapeake Bay, MD

Abstract

Previously, we reported that microplastic volatile organic compounds are present within the Chrysaora chesapeakei of Chesapeake Bay, MD. In this study, we report the presence of contaminants of emerging concern (CECs) on the hydrophobic surface of microplastic (MP) particles extracted from the C. chesapeakei, detected by Raman spectroscopy and identified by Wiley’s KnowItAll Software with IR & Raman Spectral Libraries. C. chesapeakei encounters various microplastics and emerging contaminants as it floats through the depths of the Patuxent River water column. This study identifies subsuming CECs found directly on microplastics from within C. chesapeakei in the wild using Raman spectroscopy. Among the extracted microplastics, some of the emerging contaminants found on the different microplastics were pesticides, pharmaceuticals, minerals, food derivatives, wastewater treatment chemicals, hormones, and recreational drugs. Our results represent the first of such findings in C. chesapeakei, obtained directly from the field, and indicate C. chesapeakei’s relationship with microplastics, with this species serving as a vector of emerging contaminants through the marine food web. This paper further illustrates a relationship between different types of plastics that attract dissimilar types of emerging pollutants in the same surrounding environmental conditions, underscoring the urgent need for further research to fully understand and mitigate the risks that MPs coexist with contaminants.

1. Introduction

Microplastics (MPs), a form of plastic smaller than 5 mm, are global, pervasive pollutants and contaminants of emerging concern. Distributed in the biosphere and accumulating in water bodies, MPs can be categorized as either primary or secondary. Primary MPs originate from the industrial manufacturing of personal care products, medicines, and textile products. In contrast, secondary MPs result from biological, physical, and chemical degradation processes such as natural weathering and UV radiation exposure [1,2]. Common MPs, including polyamides (PAs), polystyrenes (PSs), polyethylenes (PEs), polyvinyl chlorides (PVCs), polyethylene terephthalates (PETs), and polyacrylamide (PAM), are all composed of hydrocarbons that contribute to their electrostatic and hydrophobic properties. Plastic pollution and microplastics are major global concerns that pose pervasive threats to aquatic and terrestrial food webs, including many species commercially used for human consumption [3]. Approximately 15–51 trillion plastic particles are floating on the world’s oceans’ surfaces, representing only ~1% of the 4.8–12.7 million tons of plastics thought to enter global oceans annually [3]. Plastics are especially problematic when present in micro- and nanoparticulate forms [4]. Micro- and nanoplastics last for centuries, making them impossible to remove due to their stable properties and unrivalled threats to aquatic life. MPs’ danger to aquatic and human life lies in their large specific surface area and highly hydrophobic properties; these properties allow them to efficiently adsorb and enrich hydrophobic organic pollutants, which threaten ecosystems and humans because they are easily ingested by organisms [5,6,7,8]. Microplastics’ hydrophobicity lead to an increase in the contaminants of emerging concern (CECs) that they constantly encounter in aquatic environments.

1.1. Concerning Emerging Contaminants (CECs)

The United Nations Educational Scientific and Cultural Organization [9] defines CECs as naturally occurring or synthetic chemicals or any microorganisms that are not regulated or monitored in the environment but have adverse effects that threaten ecological and human health [9]. These chemicals include pesticides, industrial and household products, pharmaceuticals, personal care products, metals, surfactants, industrial additives, and solvents. Most of these have been known to be used, released, and then distributed into the environment, posing an ongoing threat to biota. Unfortunately, even in low quantities, CECs can cause chronic toxicity and endocrine disruption in humans and aquatic wildlife [9]. The Network of Researchers and Regulators on Energy Environmental Substances (NORMAN) defines a contaminant as any chemical, physical–chemical, radiological, or biological substance or matter that adversely affects air, water, or soil. This network further defines a contaminant of emerging concern [9]. More than 100 million chemical substances are currently registered in the Chemical Abstracts Service (CAS), and about 4000 new ones are registered every day. The European Union legislation for the Registration, Evaluation, Authorization, and Restriction of Chemicals lists 30,000–50,000 industrial chemicals in daily-use products, all of which are ultimately released into the environment [9,10].

1.2. Absorption of Emerging Contaminants by Microplastics

MPs can absorb contaminants present in environmental media because of their lipophilicity [11]. When an MP degrades into smaller plastic particles, this enhances the adsorption of contaminants because more of the MP’s surface area is exposed, and its chemical reactivity increases [12,13,14,15,16]. Factors that influence absorption include environmental conditions like weathering, sunlight, pH, long exposure times, zeta potential and hydrophobicity, which influence the kinetics of adsorption of contaminants of MPs [12,13,14,15,16]. The adsorption of contaminants occurs on MP surfaces because of MPs’ large surface area and hydrophobicity, further inducing the attachment of pollutants attached to them upon their release into the environment [12,13,14,15,16]. The adsorption of contaminants on and their desorption from MP surfaces is complicated under heterogeneous environmental conditions because of a mixture of dynamic factors such as the MP’s characteristics (e.g., composition/type, structure, binding energy, and surface properties), the release medium (e.g., pH, temperature, salinity, and ionic strength), and contamination factors (e.g., solubility, redox state, charges, and stability) [12,13,14,15,16].

Absorption and desorption kinetics differ for different types of MPs, such as polyethylene (PE), polypropylene (PP), polystyrene (PS), and polyvinyl chloride (PVC), with PE (rubbery polymer PE) having higher adsorption than the other types of MPs [17]. The absorption mechanisms of MPs involve electrostatic and hydrophobic interactions, which govern the properties of adsorbent plastic materials [18].

Recent research evidence has documented the relevant role of microplastics as vectors of contaminants, a complex aspect of the issue, as reported in [19,20,21,22,23]. This complexity adds another layer to the challenge faced. For biological contaminants, hydrological forces such as winds and currents can transport plastic-attached organisms over long distances [24].

1.3. C. chesapeakei

Similarly, C. chesapeakei is an organism whose movement is also controlled by winds and water currents [25], through which it constantly encounters the CECs of the Patuxent River Chesapeake Bay. C. chesapeakei, of the cnidaria phylum, is an invertebrate that dwells in various water depths from the bottom to the top of the Patuxent River, Chesapeake Bay, during the summer months of June through early September, when the water temperature ranges from 22 to 25 °C [25]. The stages of its growth are polymorphic and sessile in the polyp stage but mobile in the medusa stage. Stages of their development can range from the bottom of the estuary floor to a higher water depth level, depending on where their eggs can latch onto and develop. They can grow in any ocean debris as well [25]. They can live for up to six months in the wild, with incessant feeding dependent upon a sequence of chemically mediated behaviors, triggering (a) a discharge of cnidocytes by compounds usually associated with the cell membranes, mucin, and chitin of the prey, such as N-acetylated sugars; (b) a retraction of tentacles triggered by endogenous compounds to move captured prey to the mouth; and (c) the ingestion of the prey, facilitated by reversed ciliary beating in the mouth and pharynx [25,26,27,28]. Their different growth development stages in the water allow them to encounter microplastics at different water column depths [29] and in their different growth stages.

1.4. Microplastic Movements and C. chesapeakei

C. chesapeakei fares better than many other sea creatures in the eutrophic polluted waters of the Patuxent River as it does not need much oxygen to flourish and faces minimal threat by predators; therefore, it can capture contaminants with ease as it moves through the water column due to the hydrophilic bodies and the hydrophobic relations of MPs in hydrophilic environments. Microplastics move with water currents due to their light weight and specific chemical and physical properties, giving jellyfish involuntarily or voluntarily abilities to capture microplastics within their gelatinous bodies [25,29,30]. Moreover, marine pollutants induce feeding behaviors in C. chesapeakei [25,31,32,33,34]. Since microplastics cannot decompose in the Patuxent River’s water temperatures of 6.5 °C to 26 °C, they can become trapped by jellyfish through vortex pressures [25,35,36,37,38] and hydrophobic bodies containing CECs.

1.5. C. chesapeakei with Microplastics and Contaminants of Emerging Concern

This study aimed to uncover and isolate the microplastics present within the C. chesapeakei in their natural habitat; and that these species are carriers of emerging contaminants. The Raman spectroscopy, a pivotal tool in our research, meticulously and accurately identified specific copolymers of the microplastics found in C. chesapeakei jellyfish and analyzed the contaminants of emerging concern present in these microplastics.

2. Materials and Methods

2.1. Sample Collections

The Patuxent River empties into the west side of Chesapeake Bay, 143.71 km above the Virginia Capes. Commercial traffic consists chiefly of shellfish and shells and petroleum products. The river has natural depths of 8 to 9 m in the approach of 9 to over 30.5 m for 26 km upstream.

As previously reported [25], details of our successful sample collections are as follows. During the hot summers (23–29 °C) of 2021 and 2022, approximately forty C. chesapeakei specimens of various sizes were collected using a twenty-foot-handle metal strainer, and all river water was drained from the jellyfish. Sizes of C. chesapeakei specimens collected and the experimental methods used adhered to humane standards. The coordinates of the collection were as follows (Figure 1):

38°23.104 N, 076°30.025 W;

38°23.470 N, 076°29.584 W;

38°26.170 N, 076°29.386 W;

38°25.481 N, 076°29.372 W.

2.2. Contamination Prevention

All the jellyfish caught, totaling 1478 mL (measured in volume units due to the JFs’ water consistency), were distributed equally (739 mL) into two separate sterile glass jars. A small amount of 100% glycerol (0.01) was added to the samples, and they were then frozen at −20 °C until aliquots were needed for examination

Small and medium-sized C. chesapeakeis were humanely captured in the lowest amounts required at the coordinates mentioned in this study. Their medusas ranged from approximately 1.27 cm to 20.32 cm per diameter.

2.3. Raman Scattering Spectroscopy

Samples prepared for Techniques I, II, and III were measured using a micro-Raman setup based on an Olympus microscope and a Horiba JY T64000 (Horiba, Austin, TX, USA) spectrometer with liquid-nitrogen-cooled CCD. The laser probe measured 2 microns in diameter. Spectra were measured upon excitation with a 647 nm line of an Ar-Kr laser using a power of 1 mW. The original spectra contained Raman spectra superimposed against a strongly luminescent background, this luminescence originating from the dyes and defects in the plastic, which appeared due to weathering [39]. A photoluminescent background was fit using a sum of broad Lorenzian bands with a line width larger than 500 cm⁻¹ and subtracted from the spectra using LabSpec5 software. The resulting Raman scattering lines were compared to the spectra of plastics and CECs available through the Raman reference we used, Wiley’s KnowItAll Software with IR & Raman Spectral Libraries (Wiley Science Solutions, Hoboken, NJ, USA). Notably, the microplastic particles examined had to be extracted, dried, isolated and examined from within the C. chesapeakei specimens to accurately determine what may have been directly on the microplastics from within the jellyfish.

2.4. Techniques I, II, III, IV

Since no techniques had previously been established for extracting microplastics from jellyfish tissue, three different techniques were created for this study to determine microplastics’ presence in the naturally liquid gelatinous tissue. Consequently, each technique identified three different types of plastics with different absorption properties and varying levels of emerging contaminants.

The entire bodies of the jellyfish were frozen, and since jellyfish consist of 95% water, the species froze and thawed similarly to water. After defrosting when needed, the gelatinous tissue was centrifuged, as noted in the descriptions of the technique we used, resulting in a pellet containing materials obtained directly from the tissue.

In this experiment, we focused solely on extracting microplastics from the pellet; however, other substances were likely also present. Each method used a fresh, aliquoted, defrosted sample.

Technique IV did not utilize Raman Spectroscopy. Instead, it examined the tissue sample directly for minerals through a portable LED microscope.

Techniques

Technique I: A 5 mL fresh sample from collection two was centrifugated and spun at 5000 rpm with an IEC Clinical Centrifuge for 20 min in sterile glass centrifuge tubes using a modified version of the methodologies described by [40] Hildebrandt et al. 2021 and [41] Cai et al. 2021. The decanted supernatant was saved and stored in a freezer. The pellet was digested with 30% H₂O₂ at 60 °C for seven hours in a dark environment within a glass test tube, rinsed with deionized water, dried (2 h) and then observed with Raman spectroscopy on Al₂O₃ filters (Cytiva, Alvaro Obregon, Mexico City, Mexico) [Figure 2 and Figure 3].

Technique II: A fresh 2 mL thawed sample from collection two was placed in the filtration apparatus with Anodisc TM Al₂O₃ filters (Cytiva, Alvaro Obregon, Mexico City, Mexico) measuring 2.0 µm, as detailed in Medina and Taylor 2021, [42] and then observed under a Raman spectrometer [Figure 4 and Figure 5].

Technique III: A 5 mL fresh sample from collection two was used with the centrifugation procedure. The pellet digestion (again with 30% H₂O₂) time was shortened to approximately 80 °C for three hours in dark conditions; the pellet was rinsed with deionized water and dried, and then a 25 mm (2.0) µm halved nucleopore filter (Cytiva) was added to a microscopic slide, with a drop (via glass dropper) of the digested sample added to the filter paper. A cover slide was taped to the microscopic slide to secure the sample for analysis with Raman spectroscopy [Figure 6].

Technique IV: A dropper with a 0.25 mL gelatinous jellyfish tissue solution was dropped on a standard glass slide of the C. chesapeakei from the field. The samples were observed under an LED microscope (Elikliv EDM11S LCD digital microscope) with a 32GB SD card; this was a 2000× biological microscope with a digital and microbial lens, 7″ IPS display, 10 LEDs, and a resolution of 12MP, and it is Windows- and Mac OS-compatible.

3. Method I Results

Raman Spectroscopy visuals of MPs extracted from within the C. chesapeakei in the field using Techniques I–III.

3.1. Technique I Results

Figure 2. Visual image of a banana-shaped microplastic 0.0015 mm particle on a glass slide with an Al₂O₃ filter (sample obtained by Technique I). The bar in the lower right corner shows the scale, which is 4 microns. The red dot represents the laser probe, with a wavelength of 647 nm, which is focused on the studied particle.

Figure 2. Visual image of a banana-shaped microplastic 0.0015 mm particle on a glass slide with an Al₂O₃ filter (sample obtained by Technique I). The bar in the lower right corner shows the scale, which is 4 microns. The red dot represents the laser probe, with a wavelength of 647 nm, which is focused on the studied particle.

Figure 3. Lower panel: Raman spectra (red) of sample A obtained using Technique I. The upper panel shows the precise range of Raman vibrations, in cm⁻¹, of the co-polymers identified in the spectrum by the Wiley database software within the tested unknown sample, sample A. The black vertical lines match the Raman shift in cm⁻¹ and identify the position of the precise bonds and numerical spectral peaks of each copolymer within sample A.

Figure 3. Lower panel: Raman spectra (red) of sample A obtained using Technique I. The upper panel shows the precise range of Raman vibrations, in cm⁻¹, of the co-polymers identified in the spectrum by the Wiley database software within the tested unknown sample, sample A. The black vertical lines match the Raman shift in cm⁻¹ and identify the position of the precise bonds and numerical spectral peaks of each copolymer within sample A.

3.2. Technique II Results

Figure 4. Visual images of microplastic particles extracted from the jellyfish on a glass slide (sample B obtained using Technique II). The bar in the lower-right corner shows the scale, which is 4 microns. The red dot represents the laser probe, with a wavelength of 647 nm, which is focused on the approximately to 0.0012 mm particle.

Figure 4. Visual images of microplastic particles extracted from the jellyfish on a glass slide (sample B obtained using Technique II). The bar in the lower-right corner shows the scale, which is 4 microns. The red dot represents the laser probe, with a wavelength of 647 nm, which is focused on the approximately to 0.0012 mm particle.

Figure 5. Lower panel: Raman spectra (blue) of sample B obtained using Technique II. The upper panel shows the range of precise Raman vibrations that align with the Raman shift in cm⁻¹ of the polymers identified in the spectrum by the Wiley KnowItAll Edition software reference database for the tested sample. The black vertical lines match the Raman shift in cm⁻¹ and identify where the precise bonds lie in terms of the Raman shift in cm⁻¹ and in the sample, and where the spectral peaks of each copolymer are within sample B.

Figure 5. Lower panel: Raman spectra (blue) of sample B obtained using Technique II. The upper panel shows the range of precise Raman vibrations that align with the Raman shift in cm⁻¹ of the polymers identified in the spectrum by the Wiley KnowItAll Edition software reference database for the tested sample. The black vertical lines match the Raman shift in cm⁻¹ and identify where the precise bonds lie in terms of the Raman shift in cm⁻¹ and in the sample, and where the spectral peaks of each copolymer are within sample B.

3.3. Technique III Results

Figure 6. (a) Visual images of the 0.0013 mm microplastic particles on a glass filter (sample C obtained using Technique III). A bar in the lower-right corner shows the scale, which is 4 microns the laser probe used had a wavelength of 647 nm (b) Lower panel: Raman spectra (green) of sample C obtained using Method III; the upper panel shows the range of precise Raman vibrations that align with the Raman shift in cm⁻¹ of the polymers identified in the spectrum by the Wiley reference database for the tested sample. The black vertical lines match the Raman shift (cm⁻¹) and identify where the precise bonds lie in terms of the Raman shift in cm⁻¹ and in the sample, as well as where the spectral peaks of each copolymer are within the MPs on the specimen sample.

Figure 6. (a) Visual images of the 0.0013 mm microplastic particles on a glass filter (sample C obtained using Technique III). A bar in the lower-right corner shows the scale, which is 4 microns the laser probe used had a wavelength of 647 nm (b) Lower panel: Raman spectra (green) of sample C obtained using Method III; the upper panel shows the range of precise Raman vibrations that align with the Raman shift in cm⁻¹ of the polymers identified in the spectrum by the Wiley reference database for the tested sample. The black vertical lines match the Raman shift (cm⁻¹) and identify where the precise bonds lie in terms of the Raman shift in cm⁻¹ and in the sample, as well as where the spectral peaks of each copolymer are within the MPs on the specimen sample.

3.4. Technique IV Results

Rock, mineral, and crystal snowflake formations (as shown below in Figure 7), from the 0.25 mL gelatin solution on the standard glass slide on which the samples of C. chesapeakei from the field were placed, were observed under the LED microscope (Elikliv EDM11S LCD digital microscope), which had a 32GB SD card; this is a 2000× biological microscope with a digital and microbial lens, a 7″ IPS display, 10 LEDs, and a resolution of 12MP resolution that is Windows- and Mac OS-compatible.

4. Method II Results

Raman Spectroscopy identified the components of microplastics along with the concerning emerging contaminants (CECs) on each separate type of microplastic derived from Techniques I, II and III.

The results of techniques I–III indicate the type of copolymers present in each microplastic and the CECs identified on each microplastic, as detailed in the following tables according to the method used. In each table, the CECs are primarily in groups of three to five, because this is how they were identified and found on each microplastic piece by Raman Spectroscopy.

4.1. Technique I Results

The piece of MP obtained using Technique I contained the copolymers polyvinyl benzoate (PVB), polydiallyl phthalate (PDAP), polyester urethane (PUR), and polyethylene terephthalate (PET). Some CECs were identified on the 0.0015 mm microplastic piece, as can be seen in Figure 2 and described in Figure 3.

Raman spectroscopy also captured the CECs on the MP; some of these CECs are listed in

Table 1. Technique I exposes some of the concerning emerging contaminants found on the 0.0015 mm MP pieces, shown in Figure 2, extracted from within the C. chesapeakei samples. ““µ MP” in the table represents the average of the ratio of the chemicals found on the 0.0015 mm MP piece. The “ MP” column represents how many times the molecules appeared on the MP extracted.

Concerning Emerging Contaminant	Tech	µ MP	Uses	MP
Deuterium Oxide	I	0.23	microplastics, pharmeceutical additives, DNA labeling	50
Ferric phosphate tetrahydrate	I	0.26	agriculture, ceramics, and wastewater treatment	49
3-Chloro-6 hydrazinopyridazine	I	0.19	agrochemical research, synthesizing various drugs and pesticides	30
Sulfasalazine	I	0.2	considered a disease-modifying anti-rheumatic drug, pharmeceuticals	28
Sodium hexametaphophate	I	0.22	food emulsifier, softener	14
7-Hydroxcoumarin-4-acetic acid	I	0.2	pharmeucetical and biochemistry	10
Trichlomethiazide	I	0.24	pharmeceutical, diuretic	5
3,5 Dichlorobenzenesulfonyl chloride	I	0.16	used to prepare organic compounds such as medicines, dyes, and pesticides	4
5-(4-Morpholinyl)-2-nitrophenol	I	0.28	pharmaceuticals, analgesics, soluble sulfanilamides	3
Stronium titanate	I	0.11	a diamond simulant, mineral, man made	3
4-Fluoro-2-(trifluoromethyl) benzenesulfonyl chloride	I	0.13	pharmaceutical, agrochemical, or material science	2
6,7-Dihydroxycoumain-4-acetic acid	I	0.11	biological and pharmaceutical, sunscreens, and inhibits cancers cell	2
Bromopentafluoroacetone	I	0.15	pharmaceuticals, agrochemicals, and specialty chemicals	2
Clorsulon	I	0.09	veterinary medicine for the treatment of liver fluke, worms	2
Indole-3- pyruvic acid	I	0.085	in mice, reduces diarrhea, colonic inflammation, expression of certain genes.	2
p-Chlorolphenoxy acetic acid	I	0.16	synthetic pesticide	2
Phenethyl caffeiate	I	0.11	antineoplastic, anti-inflammatory antioxidant, an antiviral agent, antibacterial	2
Zinc trifluoromethanesulfonate	I	0.15	mineral	2
3,4 Difluorobenzenesulfonamide	I	0.09	synthesis of pharmaceuticals, agrochemicals, and other organic	1
1,(4-Methylphenyl)piperazine	I	0.08	pharmeceutical	1
1,4,6-Pragnatrien-3,20-dione	I	0.08	pharmeceuticals, forensics	1
2-(4-Bromophenyl)	I	0.1	flame retardant	1
4-Pregnen-11B,17,21-triol-3,20-dione	I	0.08	Hydrocortisone	1
4,6-Estradien-3,17-dione	I	0.11	estrogenic, non-aromatic steroid, minor estrogenic effects required in males	1
5-Chloro-2-methoxybenzenesulfonyl	I	0.15	pharmeceuticals	1
5-Methyl resorcinol monohydrate	I	0.15	pharmeceuticals	1
6-Chloronicotinic acid	I	0.15	agrochemicals, animal food enrichment, and food additives, pharmeceuticals	1
Calcium titanate	I	0.19	ceramics, ores of titanium	1
Clopidol	I	0.08	veterinary medicine as a coccidiostat	1
Sodium bis(trifluoromethanesulfonyl)imide	I	0.1	electrolyte in batteries and fuel cells	1
Sulfamide	I	0.15	antibiodic, nonantibiodic, pharmeceutical	1
Zirconium Silicate	I	0.13	enamels, and ceramic glazes, occurs in nature as mineral zircon	1

below.

4.2. Technique II Results

The piece of MP obtained using Technique II contained copolymers such as polyethylene 2 cyanoacrylate (PECA), poly 2-hydroxyl propyl methacrylate, poly tetramethylene terephthalate (PTT), polyvinyl benzoate (PVB), and poly diaphyll phthalate (PDAP) from Figure 4 and Figure 5. Raman spectroscopy captured the CECs on the MP, which are listed in

Table 2. Technique II exposes some of the concerning emerging contaminants found on the 0.0012 mm MP piece, shown in Figure 4 and described in Figure 5, extracted from within the C. chesapeakei samples. The “µ MP” column represents the average of the ratio of the chemicals found on the 0.0015 mm MP piece. The MP column represents how many times the molecules appeared on the MP extracted.

Concerning Emerging Contaminants	Tech	µ MP	Uses	MP
2-(bromomethyl)-1,4 bis(trifluoromethyl) benzene	II	0.11	PFAS and sometimes referred to as "forever chemicals	1
2-Acetylpyrrole	II	0.13	It is used as a food additive in items such as cocoa, rum, brandy, and caramel	1
2-Aminopyridine-4-carboxylic acid	II	0.16	agrochemicals and pharmaceuticals.	1
2-Bromo-4-methoxy-6-nitrophenol	II	0.14	chemical properties, making it a versatile intermediate in various industrial applications.	1
2-Deoxyctidine-5-monophosphate	II	0.28	synthesis of DNA and RNA and commonly produced by bacteria E.coli geneses in the wild	36
2-Methyl-4-nirtoaniline	II	0.35	aniline compound used in the synthesis of dyes and pigments	36
4-[(E)-1,3-Thiazol-2-uldiazenyl]-1,3-benzenediol	II	0.14	pharmaceutical affiliate	1
4-Amino-2-nitrophenol	II	0.16	pH indicator, synthesis of dyes, wood preservatives, photographic developers, and explosives	1
4-Nitrophenylhydrazine	II	0.32	exhibits potential anticancer activity and acts as a derivatizing agent for carbonyl compounds	3
4,5-Dichloro-2-nitroaniline	II	0.15	used in industrial settings and as a component in materials such as flooring, furniture, and toys	1
5-Chloro-4-methyl-2-nitroaniline	II	0.16	starting material for the production of dyes, pharmaceuticals, and some pesticides	2
6-Mercaptopurine monohydrate	II	0.14	used to treat acute lymphoblastic or lymphocytic leukemia and used to treat cancers	1
6-Methyl-2-nitro-3-pyrindinol	II	0.2	a chemical intermediate in the synthesis related to pharmaceuticals and materials science	1
Barium Anhydrous	II	0.13	manufacturing, analytical chemistry, and testing	1
Barium Pyrovanadate	II	0.17	hard, ductile transition metal, primarily used as a steel additive	1
Benzimidazole-5-6-dicarboxylic acid	II	0.15	agrochemical, pharmaceutical, and dyestuff field	1
Cromophtal Violet B	II	0.1	organic pigment for coloring PVC, PO, PS, PET, rubber, PUR and PP. Also used in fibers.	1
Cyclocreatine	II	0.08	antiviral, antidiabetic, protect tissues from hypoxic, ischemic, neurodegenerative or muscle damage	1
Diethylenetriamine pentaacetic acid	II	0.09	inactive ingredients for drug products by the FDA, DPTA was developed by the pharmaceutical company	1
Dumorterite	II	0.23	minerals used in the manufacture of high grade porcelain.	39
EDTA. dipotassium salt dihydrate	II	0.13	textiles	1
Furazolidone	II	0.12	oral treatment against bacterial and protozoal infections	1
Methyl 3-isothiocyanatopropionate	II	0.09	chemical intermediate in the development of agrochemicals and pharmaceuticals,	1
Methyl difluoroacetate	II	0.08	fruity-smelling liquid used as an intermediate in the synthesis of pharmaceuticals and agrochemicals	1
p-(p-Phenylene terephthalamide)	II	0.12	reinforcing composites in rubber industries, such as hoses, conveyor belts, tires, and rubber tracks	1
Poly(acetal) reinforced	II	0.13	polyformaldehyde, is an engineering thermoplastic used in precision parts	1
Poly(dimethylolurea)	II	0.08	Antimicrobial characteristics, wood treatment, textile finishing, tanning and photographic developers	1
Propyleneimine	II	0.1	chemical is used in the paper, textile, rubber and pharmaceutical industries, paint	1
Salvinorin A	II	0.13	hallucinogen, unique pharmacological effects on opioid receptors,	1
Thallium acetate	II	0.14	selective agent against gram-negative bacteria in selective media for the detection of mollicute	1
Triacetin	II	0.11	cosmetic biocide (most often as a fungicide), plasticizer, food additive (as a flavoring agent and adjuvant),	1
Tungstic acid	II	0.1	mordant and a dye in textiles	1

below. A graph of CECs from Technique II on the 0.0012 mm MP piece from the field is also presented.

4.3. Technique III Results

The piece of MP obtained using Technique III contained the copolymers polyvinyl pyrrolidine (PVP), polyacrylamide (PA), polyvinyl methyl ketone (PVK), and polymethylene urea (PMU) from Figure 6a,b. Raman spectroscopy captured the CECs on the 0.0013 MP piece, some of which are listed in

Table 3. Technique III exposes some of the concerning emerging contaminants found on the 0.0013 mm MP piece, shown in Figure 6a and described in 6b, extracted from within the C. chesapeakei samples. The “µ MP” column represents the average of the ratio of the chemicals found on the 0.0013 mm MP piece. The “ MP” column represents how many times the molecules appeared on the MP extracted.

Concerning Emerging Contaminants	Tech	µ	Uses
1,1,2-Trichlorotrifluoroethane	III	0.11	refrigerant, a dry cleaning solvent, an extraction solvent for petroleum hydrocarbons, oils, and greases	1
Adamite	III	0.13	zinc arsenate hydroxide mineral,	2
Aluminum Sulfate	III	0.14	coagulating agent purification of drinking water, wastewater treatment plants, paper manufacturing	1
Amblygonite	III	0.08	phosphate mineral with lithium and fluorine, often found in granite pegmatite	1
Ammonium magnesium chloride hexaydrate	III	0.15	dehydrated to form anhydrous magnesium chloride, which is used in various industrial processes	1
Ammonium sodium phosphate dibasic tetra	III	0.15	employed in the food industry as a food additive,	1
Aniline sulfate	III	0.12	manufacturing of polyurethan plastics, paracetamol, acetaminophen, Tylenol, pesticide and fungicide.	16
Beryl	III	0.11	mineral composed of beryllium aluminium silicate	1
Beryllonite	III	0.09	phosphate mineral	1
Carundum(sapphire)	III	0.09	mineral	1
Diopse	III	0.08	rare mineral occurring as emerald green or blue-green crystals made of hydrated silicate of copper	1
Enstatite	III	0.11	a mineral composed of Magnesium	3
Epidote	III	0.13	aluminum-iron mineral that can be found in either crystal or stone form	1
Euclase	III	0.12	beryllium aluminium hydroxide silicate mineral	2
Forsterite	III	0.15	magnesium-rich olivine mineral that forms in igneous and metamorphic rocks	2
Fuamaryl chloride	III	0.13	chemical intermediate used in the production of pharmaceuticals, dyestuffs, and insecticides	37
Lead sulfate	III	0.16	a sulfate of lead that can occur naturally as the mineral anglesite, and often in lead-acid batteries	2
Methylbenzoylecgonine HCl	III	0.09	major metabolite of cocaine	1
Phenakite	III	0.18	mineral made of of beryllium orthosilicate	1
Phosphoric acid solution	III	0.16	produced from phosphate rock	1
Poly(chlorotrifluoroethylene	III	0.08	is a thermoplastic chlorofluoropolymer	1
Pyromorphite	III	0.35	mineral species composed of lead chlorophosphate	42
Rutile	III	0.15	an oxide mineral composed of titanium dioxide (TiO2), the most common natural form of TiO2.	3
Seripierite	III	0.23	rare, sky-blue coloured hydrated sulfate mineral, often found as a post-mining product	2
Titanite	III	0.11	titanium mineral accessaory to granitic and calcium rich metamorphic rock	2
Zinc borate	III	0.15	flame retardant in plastics and cellulose fibers, paper, rubbers and textiles, paints, adhesives, pigments	7

below.

5. Discussion

Marine microplastic litter from the Patuxent River, Chesapeake Bay, is a primary concern since it accumulates highly hydrophobic organic and inorganic pollutants in a short time, making it an ideal source of attraction for the gelatinous body of C. chesapeakei [43,44,45,46]. Consequently, the short accumulation period gives ample time for the C. chesapeakei, during its brief life cycle of no longer than six months in the wild, to accumulate microplastics as it floats through the water column [47,48].

C. chesapeakei’s contact with microplastics and CECs is evident [25], as both the CECs and microplastics of Chesapeake Bay exist in significant numbers [49] in the Patuxent River, MD. Even though the distinct spatial distribution of CECs in water, sediment, and biota remains unknown, the CECs of the Chesapeake Bay, the largest estuary in the United States [49,50], have been previously measured. For example, in the surface water of rural areas of Chesapeake Bay [51,52,53,54], antibiotics, pharmaceuticals and hormones have been detected [50], potentially stemming from animal feeding operations (AFOs) [55] or shallow groundwater discharges. While the occurrence of antibiotics, hormones, pharmaceuticals and personal care products has been reported in more populated areas of the Bay, many of them match what was demonstrated on the microplastics (Figure 2, Figure 4 and Figure 6) [50,53,56,57,58] removed from the Chrysaora Chesapeakei samples. Previous studies have focused on at least 43 antibiotics, nine hormones, 11 UV filters, and various sucralose concentrations in matched water, sediment, and oyster samples from 58 sites distributed across the Maryland section of Chesapeake Bay, including the Patuxent River area. In conjunction, 64 CECs and 58 sites established confirmed contaminant profiles in regions differentially influenced by WWTP septic systems. Patuxent River has some of the most significant distributions of wastewater treatment plants in the Chesapeake Bay watershed that input CECs [51,52,53,54]. Consequently, the MPs extracted from the C. chesapeakeis samples contained CECs such as dumortierite (stone) (Table 2), fumaryl chloride (pharmaceutical) (Table 3), 2-deoxycytidine-5-monophosphate (DNA) (Table 2), sulfasalazine (antibiotic) (Table I), 3-Chloro-6-hydrazinopyridazine (pesticide) (Table 3), and 2-acetylpyrrole (sugar flavoring); hence, similar or the exact same substances have been verified by previous researchers who have focused only on Chesapeake Bay water, fish and oyster studies. Further studies would be beneficial in determining all the origination points of the CECs found in the microplastics extracted from C. chesapeakei.

The presence of CECs on MPs from the C. chesapeakei samples and the subsequent validation of previous research on sorption and desorption levels demonstrate the complexity of the relationship between contaminants and microplastics. The unique molecular structure of contaminants, leading to their differential absorption and desorption on each microplastic, is demonstrated in three different types of plastic (Figure 2, Figure 4 and Figure 6). Different CECs present on different compositions of microplastics found (Table 1, Table 2 and Table 3) in the same jellyfish species and collected at similar times expose the intricate nature of our research, clarifying that different types of MPs can carry different CECs [17,18]. Isolating and examining microplastic particles from the C. chesapeakei samples was crucial in our research. This meticulous process provided an accurate determination of certain substances’ direct presence on the microplastics extracted from the jellyfish.

Recent studies have uncovered a complexity in microplastic (MP) analysis, demonstrating that rock complexes exist in MPs [59]. The MPs extracted from the C. chesapeakei samples align with these findings. The MPs in the C. chesapeakei samples collected from the field contained certain rocks, gems, and stones, as identified by Raman spectroscopy (Table 1, Table 2 and Table 3). Further, as shown in Figure 7 when the C. chesapeakei solution did not show distinct microplastics, rock-like crystals were solely visible under an LED microscope [36], further solidifying the reliability of our findings.

Specific hydrostatic, polar, and hydrogen bonding helps to define microplastics under various conditions. Additionally, C. chesapeakei can survive under hypoxia levels (1.5 mg/li), providing a conducive environment for microplastic pollutants to enter the jellyfish, since they are surrounded by certain pollutants at low-oxygen levels [25,60]. The salinity levels of the Patuxent River also appear to be prime for pollutants to thrive and adhere to microplastic sites, as higher salinity levels (10–15 PSU) induce microplastic adhesion to CECs [60]. Furthermore, the constant alkaline pH level of the Patuxent River, ranging from pH 8 to 8.50, [60] enhances the absorption of CECs by microplastics.

A challenging part of this research is that CECs need to be included in routine environmental monitoring programs and should be candidates for future legislation due to their adverse effects and/or persistence [9,10]. CECs are substances with a lifecycle, a behavior, and (eco)toxicological effects that are not entirely understood [9,10]. Further, CECs in the marine environment appear infinite [9,10]. Many are unlikely to become CECs, but the need for better regulation is evident [9,10]. Numerous international environmental agencies and regulators have compiled individual lists of chemicals and substances they regard as concerning [9,10]. Still, the need for a standard list is a challenge to meet.

6. Conclusions

Microplastics trapped in the gelatinous surface of jellyfish (C. chesapeakei) caught from the Chesapeake River (Maryland, USA) were isolated using three different extraction techniques. Each technique was carefully selected and applied, as it was necessary to identify the distinct influence of CECs’ intermolecular forces on its absorption by various types of microplastics, such as polyvinyl chloride (PVC), polyethylene two cyanoacrylate (PECA), polyethylene terephthalate (PET), and polyacrylamide (PA). The chemical nature or the types of isolated microplastics, the presence of chemical additives and heavy metals (coloring agents) within the MPs, and the absorbed contaminates from the contaminated Chesapeake River water on the hydrophobic surface of MPs were further analyzed by Raman spectroscopy, followed by analysis using the Wiley KnowItAll software database, as some of the results reveal in Table 1, Table 2 and Table 3. Specific types of plastics within the C. chesapeakei jellyfish in the wild were found to carry different types of CECs on the hydrophobic surfaces of the MP. The absorption capacity of the hydrophobic surface significantly enhances confidence in the understanding that certain types of microplastics carry certain CECs better than other types of MPs. Moreover, this adds to the importance of MPs, in that they can act as vectors throughout the aquatic food web carrying toxic substances.

The recent literature has confirmed that the toxicity of MPs in the human body occurs at the cellular level and that their accumulation may damage the organ and organ system, which may even lead to death [61,62,63,64]. Accumulation of MPs itself may cause inflammation, oxidative stress, and disruption of nerve impulses, and it may alter the gut microbiota. On the other hand, additives such as BPA, phthalates, etc., are endocrine disruptors that might alter the body’s hormonal regulation. Also, heavy metals such as Pb, Cr, and so on in the coloring of plastics may act as enzymatic inhibitors. The most serious concerns are the hydrophobic surface of MPs, which traps the insoluble chemical pollutants, such as PCB, DDT from the environmentally toxic water. Recent reports state that MPs are carriers of antibiotic-resistant genes and promote long-range transport [65]. Moreover, in a 2019 study, it was found that United States citizens consume 39,000–52,000 MP particles each year from food and water [66]. The sheer volume of microplastics in our environment poses a significant threat to the human immune system. Further research on the effect of microplastics as vectors of CECs and their potential harm to human tissues and marine life through the continuous bioaccumulation of toxicants, carcinogens, mutagens, and teratogens in the food web is imperative. This underscores the fact that microplastics are a silent but grave threat to all living organisms.