Enhancement of a Simple, Economic and Eco-Friendly Analytical Approach for the Extraction and Determination of Endocrine Disruptors from Plastics in Shrimp

Abstract

The economic significance of the shrimp industry relies heavily on the comprehensive utilization of all of the shrimp’s parts. However, this importance is often threatened by common challenges such as disease and pollution, caused by prominent contaminants that are capable of exerting adverse effects either directly as physical pollutants or indirectly through the incorporation of additives or adsorbed chemicals. Among these substances are endocrine disruptors, which pose risks to both wildlife and human populations. In this study, 11 endocrine-disrupting compounds were determined (3 bisphenols, 3 phthalates, 3 pesticides, and 2 nonylphenols) through the development of a cost-effective, greener and cost-friendly method based on solid-phase matrix dispersion (MSPD) with high-performance liquid chromatography coupled with diode array detection (HPLC-DAD). Determinations were performed on different parts of the shrimp: the cephalothorax, abdomen, intestine and shell. Several variables were optimized in the extraction, separation and detection phases, resulting in average recoveries of about 90%. The limit of detection (LOD) varies depending on the analyte and matrix. At concentrations of 1 mg/kg in the cephalothorax+shells and 1.25 mg/kg in the abdomen+intestine, all compounds were detected, except for nonylphenols. The developed method has allowed the simultaneous determination of 11 endocrine disruptors in different parts of the shrimp samples. Furthermore, the MSPD has been demonstrated to be an efficacious, selective, and streamlined sample extraction method, eliminating the necessity for pretreatment steps such as centrifugation and filtration, as well as the use of large volumes of solvents.

1. Introduction

The economic importance of marine resources is beyond doubt, and in the case of shrimp, the global demand for different species makes it a crustacean of incalculable value. Shrimp for consumption or commercialization can be obtained in two ways, including fishing or aquaculture, the latter being widely used in Asian countries and becoming one of the pillars of their economies [1,2]. Worldwide, shrimp is one of the most valued species in aquaculture. Its production has experienced a significant increase, reaching 5.6 million tons in 2023 [3], from less than 75,000 tons in 1980 [4]. It is considered that the head and shell represent approximately 50–60% of the total weight of the shrimp [5]. Therefore, one of the main factors contributing to the shrimp’s importance is the utilization of all parts of the shrimp, whether for direct consumption or for the production of bioproducts (chitin, carotenoid pigments, proteins, etc.) [6,7]. The white shrimp stands out as the species of greatest commercial importance, in terms of both catch and economic value, along the Mediterranean coasts of Spain and Italy, and to a lesser extent, France. A tiny fraction of the production comes from local fisheries or the coastal trawl fleet. However, most of the shrimp sold in supermarkets is caught in the southern Gulf of Guinea by the deep-sea trawl fleet, where this stock is fully exploited or overexploited. A similar situation is observed in the fishing grounds of Northwest Africa, where white shrimp stocks appear to be heavily exploited or overexploited.

There are a number of issues that can impact both shrimp production and its economic value, including diseases and contamination by compounds that are harmful to human health. One of the contaminants in the oceans that requires urgent attention is plastics, particularly micro- and nanoplastics, due to their extensive presence in the marine environment. The prevalence of marine plastic pollution has been on the rise over recent years, with the greatest concentrations observed in the water column. This has led to an increased uptake of plastic by marine organisms, which can enter the trophic chain [8]. There is a potential for plastics to accumulate in the human body, which could pose a risk to human health [9,10]. Plastics contain additives that improve their functionality by modifying their physical and chemical characteristics, affecting their properties and manufacturing costs [11]. These additives may represent new routes of exposure to chemical compounds, particularly when ingested. The global plastic additives market was valued at $48.41 billion in 2020. It is expected to increase from $51.04 billion in 2021 to $75.20 billion in 2028, at a compound annual growth rate of 5.7% over the period 2021–2028 [12].

Conversely, the documented capacity of microplastics to adsorb a range of contaminants underscores their role as carriers. These carriers can facilitate the transport of chemical contaminants over long distances or their transfer to different organisms [13,14,15]. Many of the plastic additives and some of the adsorbed compounds are of particular concern because they are endocrine-disrupting chemicals (EDCs), which can alter the human endocrine system and affect health. The presence of EDCs in shellfish is contingent upon a multitude of variables, including the source and quantity of EDCs entering the aquatic environment, the physicochemical properties of the EDCs and their accumulation within the food chain [16]. Many of these compounds are commonly present in the marine environment due to industrial, agricultural and urban discharges that release these substances into water bodies. Even compounds that are banned in many countries, such as DDT and its derivates, appear in various environmental compartments, including marine waters. In addition, shrimp, like other aquatic organisms, can bioaccumulate these compounds in their tissues when exposed to them in their habitat. EDCs found in shrimp include phthalates [17,18], pesticides [19,20,21], nonylphenols [22] and bisphenols [23,24,25,26]. The analysis of these compounds in shrimps is typically conducted using chromatographic methods coupled with mass spectrometry [17], with a previous pretreatment step to extract and clean the samples. Among these pretreatments, classic solid-phase extraction (SPE) [22,24] and QuEChERS [17,27] are the most commonly used methods. Thus, Gu et al. developed a method for the determination of trace endocrine-disrupting chemicals, including five phthalate esters, five monoalkyl phthalate esters, four alkylphenols, and bisphenol A, in seafood. The method was developed using ultrasound-assisted extraction (UAE) in conjunction with SPE and was employed as a simultaneous extraction and purification technique for these compounds. Both liquid chromatography tandem mass spectrometry (LC-MS/MS) and gas chromatography tandem mass spectrometry (GC-MS/MS) were utilized for analysis [28]. Additionally, Azzouz et al. conducted an investigation to determine the presence of bisphenol A, two alkylphenols and other compounds in seafood samples from Europe and North Africa. The investigation employed UAE followed by continuous SPE prior to GC-MS analysis [22]. In recent years, there has been a demand for more sustainable methods for analytical extraction; among these methods, matrix solid-phase dispersion (MSPD) deserves to be highlighted due to its ease of performance, the wide range of matrices and analytes that can be analyzed, and its low use of consumables [29]. However, most of the existing methods in the literature are technically complex and expensive, limiting their applicability in laboratories with limited resources.

This work is focused on the development of an alternative, simple, greener and cost-effective method for the extraction, separation and detection of 11 EDCs (8 plastic additives: bisphenols A, F and S (BPA, BPF and BPS), di(2-ethylhexyl) phthalate (DHEP), butylbenzyl phthalate (BBP), diethyl phthalate (DEP), nonylphenol (NP), nonylphenol-9 (NP-9) and 3 pesticides as potential contaminants that may be adsorbed by plastics: dichlorodiphenyltrichloroethane (DDT) and its metabolites, dichlorodiphenyldichloroethane (DDD) and dichlorodiphenyldichloroethylene (DDE)) in shrimp samples. Figure 1 depicts the chemical structures of the selected analytes.

The method was applied to the four principal parts of the shrimp: the head or cephalotorax (CT), the abdomen (AB), the intestine (IN) and the shell (SH).

The combination of the different parts is driven by their distinct culinary applications. Typically, CT-SH (cephalothorax and shell) is not consumed directly, as it is primarily used for broths and soups to impart flavor. By contrast, the components that constitute AB (abdomen) and IN (internal organs) are consumed together in various dishes. Given the varied uses and consumption patterns of shrimp, it is important to determine the contaminant content in the different parts to ensure food safety and to understand the potential exposure risks to consumers. This information is necessary for developing targeted strategies to minimize contaminant levels in the parts of shrimp most consumed.

To this end, a matrix solid phase dispersion (MSPD) method was developed and validated, which permits several steps of sample preparation to be performed simultaneously. In this method, the sample is uniformly distributed in a solid medium, facilitating total sample fragmentation and dispersion into extremely small particles. This process results in an increased surface area, which is beneficial for the subsequent extraction of compounds. The MSPD method typically uses solvent volumes on the order of 95% less than classical methods and requires approximately 90% less time [30], making the detection of EDCs more accessible and efficient.

2. Materials and Methods

2.1. Reagents and Materials

The solvents methanol (MeOH), acetonitrile (ACN) and dichloromethane (CH₂Cl₂) (HPLC grade-purity) were purchased from Scharlab (Madrid, Spain). The analytical standards of nonylphenol (NP) (purity ≥ 99%), 9-n-nonylphenol (NP-9) (purity ≥ 99%), bisphenol S (BPS, purity ≥ 98%), bisphenol F (BPF, purity ≥ 98%), bisphenol A (BPA, purity ≥ 99.9%), di(2-ethylhexyl) phthalate (DEHP, purity ≥ 99.5%), dibutyl phthalate (DBP, purity ≥ 99%) and diethyl phthalate (DEP, purity ≥ 99%) were obtained from Sigma Aldrich (Madrid, Spain).

Individual standard stock solutions of the selected compounds, with the exception of pesticides, were prepared in methanol at a concentration of 1000 mg/L and maintained in darkness at 4 °C until use. Daily working standard solutions were prepared by suitable dilution with a methanol/water mixture (85:15, v/v). Ultrapure water was obtained using a Milli-Q water system (Merck Millipore, Madrid, Spain), which has a resistivity of 18 MΩ·cm and a temperature of 25 °C. Florisil (60–100 mesh) was procured from Acros Organics (Madrid, Spain), while sodium sulfate anhydrous (Na₂SO₄, purity ≥ 99.9%) and washed sea sand (0.25–0.30 mm) were sourced from Panreac (Barcelona, Spain).

2.2. Sampling and Preliminary Treatment

Fresh white shrimp samples originating from the Atlantic Ocean (Huelva, Spain) were procured from a local market (Madrid, Spain). A total of 70 individuals were collected, with a size range between 6.5 and 8.5 cm and weights between 9 and 11 g. These were separated into two batches of similar weight with 35 individuals in each, and stored at 5 °C in glass jars with metal lids. To evaluate the levels of contamination in which the studied analytes could be present in the different parts of the shrimp, samples from the first batch (Batch 1) were dissected and divided into four distinct groups as follows: the cephalotorax (CT), intestine (IN), abdomen without intestine (AB) and shell (SH). A second batch of samples was established (Batch 2) for the purpose of separating the parts of the shrimp that are directly consumed, such as the abdomen and the intestine, from those that are not, such as the cephalothorax or the shells, which are typically utilized in the preparation of broths or soups. Thus, the samples of Batch 2 were dissected and divided into two groups as follows: cephalotorax + shell (CTSH), and abdomen with intestine (ABIN). The combination of the different parts is driven by their distinct culinary applications. Typically, CTSH (cephalothorax and shell) is not consumed directly, as it is primarily used for broths and soups to impart flavor. By contrast, the components that constitute AB (abdomen) and IN (intestine) are consumed together. Given the varied uses and consumption patterns of shrimp, it is important to determine the contaminant content in the different parts to ensure food safety and to understand the potential exposure risks to consumers. This information is necessary for developing targeted strategies to minimize contaminant levels in the parts of shrimp most consumed. The different groups with the parts of the shrimps were then washed with ultrapure water and stored at −18 °C until analysis. In Batch 1, the cephalotorax was separated from the abdomen by removing the shell and intestine, making a slight longitudinal incision in the dorsal area of the abdomen. In Batch 2, the cephalotorax and shells were considered together according to their use, and the intestines were not removed from the abdomen. The separation of shrimp parts was performed on a metal tray using a cutting blade and tweezers. Mechanical crushing was performed with a metal hand blender in each jar to avoid additional plastic contamination, except for the intestine matrix, which was crushed and homogenized in a glass mortar due to its small volume.

During the shell pretreatment process, achieving complete crushing proved challenging due to the hardness of the shells, their low moisture content, and their relatively small mass compared to CT or AB. Initially, some parts remained uncrushed because the mixer blade could not reach certain areas of the sample. To enhance the homogeneity of the sample, it was frozen and subsequently ground in a glass mortar.

2.3. MSPD Extraction Procedure

In a typical MSPD procedure, samples are blended with a sorbent to obtain a homogeneous mixture, which is then transferred and packed into an extraction cartridge. Subsequently, solvent is passed through the cartridge to perform washing and elution steps for extracting and isolating analytes from the matrix. In this work, a modification of a previously developed procedure [31] was employed for the extraction of the analytes from spiked and blank shrimp samples. The optimized procedure involved spiking a previously homogenized 0.1 g shrimp sample with 100 μL of standard solution. Subsequently, the spiked shrimp sample was mixed with 0.5 g of Florisil, 0.5 g of Na₂SO₄ (purity ≥ 99%) and 0.2 g of washed sea sand for 10 min in a glass mortar. Once blended and homogenized, the mixture was placed within a glass column, with a plug of glass wool positioned at the base and a flow regulator installed. The analytes were eluted with a 3 × 1 mL and 2 × 3 mL sequence of methanol, regulating the flow rate with a plastic key to maximize drag. The collected extracts were subjected to filtration and subsequent evaporation under a nitrogen stream. The dry residue was redissolved in 400 μL of methanol:water (85:15) for further high-performance liquid chromatography (HPLC) analysis. The procedure for preparing the blank samples was identical, with the exception that the spiked analytes were not included in the MSPD mixture.

2.4. HPLC-DAD Conditions

The analytes underwent chromatographic separation using a liquid chromatograph from Agilent Technologies (model 1200 series, Waldbronn Germany). This device was equipped with a quaternary pump, an on-line degasser, a photo-diode array detector (DAD) and an autosampler. The separation was conducted on an ACE^® C18-PFP (ACE-1210-1546) HPLC column (150 × 4.6 mm, 5 μm), sourced from Symta (Madrid, Spain). The mobile phase was a solution of Milli-Q water (referred to as solvent A) and acetonitrile (referred to as solvent B). The gradient program commenced at 0 min and proceeded as follows: from 0 to 30 min, the percentage of solvent B increased from 45% to 80%; from 30 to 31 min, it increased from 80% to 100%; and it remained at 100% for the subsequent 7 min. A flow rate of 0.8 mL per minute was established. Subsequently, the column was reconditioned to 45% solvent B and maintained under isocratic conditions for 10 min. The sample injection volume was 20 μL. All compounds were successfully separated within a span of 38 min. To achieve optimal sensitivity, the peak areas were quantified by selecting the most appropriate detection wavelength for each compound. Consequently, all analytes were quantified at 210 nm. For the purposes of quantification, external calibration and peak area measurements were employed.

3. Results and Discussion

3.1. Optimization of MSPD Procedure

An MSPD method was employed to extract the analytes from shrimp solid samples. In this extraction method, samples can be directly mixed with a selected sorbent to obtain a homogeneous mixture, which is subsequently transferred to an SPE cartridge or column and eluted with the optimized solvent. Compared to the classical solvent extraction method, the MSPD eliminates the steps of repeated centrifugation and/or filtration, as well as procedures for re-extraction. This is unlike SPE, which typically involves a separate solvent extraction step to prepare solid samples for loading into an SPE cartridge. In addition, MSPD does not require this additional solvent-extraction step. These differences result in significant reductions in solvent consumption and the required manipulation time for sample preparation. Furthermore, in this study, sea sand has been employed as a more economical and environmentally friendly alternative solid support for the disrupting sample, replacing commonly used materials based on silica. As a result, we can assert that the MSPD methodology is more in line with green chemistry principles than other traditional extraction methods.

The applied MSPD procedure is based on a method previously developed by Cañadas et al. [31] for the extraction of phthalates, bisphenols, alkylphenols and organochlorine pesticides from raw mussels. A solid support consisting of Florisil and washed sea sand as sorbent and dispersant, respectively, was used in combination with Na₂SO₄ as a drying agent to disrupt and disperse the sample. After thorough mixing and homogenization in a glass mortar, the sample was transferred to an empty glass column. The nature of the elution solvent is a critical step in the process, as it determines the efficiency with which target analytes can be desorbed. The recovery rate is calculated by comparing the concentration of the spiked sample with the added concentration. This relationship is expressed as a percentage by multiplying the result by 100. Figure 2 illustrates the recoveries observed in distinct matrices when volumes of 3 × 1 mL and 2 × 3 mL of acetonitrile were used as the elution solvent for spiked matrices of cephalotorax–shell (CTSH) and abdomen–intestine (ABIN).

Initially, acetonitrile was used as the elution solvent. Due to the lower recoveries obtained with this in the case of the ABIN matrix, the procedure was modified, and dichloromethane and methanol were also evaluated as eluent solvents for the extraction of the analytes, using a volume of 3 × 1 mL and 2 × 3 mL in both cases. As can be observed in Figure 3, the percentage recovery for all analytes, except for DEHP, were significantly better (82–105%) when using methanol than in the case of dichloromethane and acetonitrile. To optimize the volume of solvent required for effective extraction of the analytes, different volumes (6, 9, 12 and 15 mL) of the optimal elution solvent, methanol, were tested. The best results were obtained with 9 mL. In the case of a 12 mL volume, as well as that of 15 mL, the results did not show significant improvement. Additionally, this volume led to increased analysis time and additional solvent consumption. Therefore, it is concluded that the optimal volume for elution of all analytes was 9 mL, added in two fractions of 3 × 1 mL and 2 × 3 mL.

3.2. HPLC-DAD Analysis

To obtain the most accurate analytical signal for each compound, the absorbance at different wavelengths, 210, 230, 250, 254 and 280 nm, was evaluated with the diode array detector (DAD). The largest areas for all analytes were generally observed at 210 nm, and thus this wavelength was selected as the optimal one. Figure 4 depicts the chromatogram resulting from the separation of a standard mixture of the eleven EDCs using HPLC-DAD under optimal conditions. The separation of these compounds was achieved within 35 min. Regarding chromatographic separation, it is evident that NP-9 requires optimization. Its peak is notably flat and wide, rendering it undetectable in the selected matrices.

Chromatographic analysis of the spiked and unspiked samples was performed in the Batch 1 and Batch 2 matrices. Figure 5 and Figure 6 show the chromatograms corresponding to individual Batch 1 matrices and grouped Batch 2 matrices spiked at 30 mg/L for nonylphenols and 10 mg/L for the other nine analytes. The chromatograms exhibit good resolution for all analytes, except for nonylphenols, which present difficulties for the quantification of NP, rendering NP-9 quantification impossible. The sample extracts were generally clean, with no significant presence of interfering substances. However, increased background noise was observed in the latter part of the chromatogram, potentially affecting the detection of late-eluting analytes. Despite this, the retention times of the analytes of interest were free from observable peaks attributable to matrix interferents or co-extracted compounds.

3.3. Validation of the Method with Commercially Spiked Shrimp Samples

This method was validated in terms of linearity, precision, recovery and limits of detection (LODs) and quantification (LOQs), using spiked shrimp samples. Calibration curves were constructed by spiking shrimp samples in the range of 5 × 10⁻⁴–40 mg/kg for all the analytes except for nonylphenols, where the spiking range was 25 × 10⁻⁴–120 mg/kg. The results demonstrated satisfactory linearity for all analytes, with the determination coefficients being acceptable for most of the compounds under study.

The precision of the method was evaluated in terms of repeatability (intra-day precision) and reproducibility (inter-day precision), with the results expressed as relative standard deviation (RSDrt and RSDrd, respectively). The repeatability and reproducibility were determined with three replicates at different concentrations, analyzed on the same day and on three consecutive days, respectively.

The observed RSD was compared with the level of precision predicted by the Horwitz equation. The Horwitz equation is RSD_Horwitz = 2^{(1 − 0.5logC)}, where C is the concentration of the analyte expressed as a mass fraction and RSD_Horwitz is the predicted relative standard deviation. The HorRat or Horwitz ratio, HorRat = RSD/RSD_Horwitz was calculated and used as an acceptance parameter for precision. According to the repeatability conditions, the accepted values are between 0.3 and 1.3. Conversely, under the reproducibility conditions, the values are between 0.5 and 2.

Table 1. Relative standard deviation (RSD, %), relative standard deviation by RDS Horwitz (%Horwitz) and Horwitz ratio (HorRat) in the mixture of cephalothorax + shell (CTSH) samples spiked at different concentrations.

CTSH	1 ppm		1.25 ppm		2 ppm		2.5 ppm
	%RSDHorwitz 16.00		%RSDHorwitz 15.471		%RSDHorwitz 14.415		%RSDHorwitz 13.939
	%RSD	HorRat	%RSD	HorRat	%RSD	HorRat	%RSD	HorRat
BPS	3.794	0.237	2.408	0.156	2.099	0.146	3.211	0.230
BPF	1.773	0.111	2.496	0.161	1.044	0.072	2.231	0.160
BPA	0.414	0.026	1.855	0.120	1.711	0.119	3.169	0.227
DEP	5.34	0.321	6.857	0.443	3.511	0.244	3.522	0.253
BBP	1.423	0.839	11.413	0.738	30.998	2.150	3.653	0.262
DDD	5.716	0.357	18.686	1.208	52.032	3.610	50.991	3.658
DDT	2.251	0.141	3.400	0.220	0.664	0.046	3.650	0.262
DDE	7.260	0.454	1.268	0.082	2.000	0.139	4.869	0.349
DHEP	2.570	0.161	5.784	0.374	0.762	0.053	43.424	3.115

presents the values of the Horwitz ratio in CTSH samples spiked at different concentrations. Nonylphenols were excluded from this analysis, as they were not detected at the specified concentration range. In the CTSH samples, some values exceeded 2 as follows: BBD and DDD at 2 ppm, and DDD and DEHP at 2.5 ppm.

Table 2. Relative standard deviation (RSD, %), relative standard deviation by RDS Horwitz (%Horwitz) and Horwitz ratio (HorRat) in the mixture of abdomen + intestine (ABIN) samples spiked at different concentrations.

ABIN	1.25 ppm		2 ppm		2.5 ppm
	%RSDHorwitz 15.471		%RSDHorwitz 14.415		%RSDHorwitz 13.939
	%RSD	HorRat	%RSD	HorRat	%RSD	HorRat
BPS	7.538	0.487	1.295	0.090	5.203	0.373
BPF	3.585	0.232	0.576	0.040	1.054	0.076
BPA	3.031	0,196	1.906	0.132	3.127	0.224
DEP	8.460	0.547	2.520	0.175	0.841	0.060
BBP	4.218	0.273	0.317	0.022	4.423	0.317
DDD	1.275	0.082	0.725	0.050	13.760	0.987
DDT	3.351	0.217	1.574	0.109	2.424	0.174
DDE	1.697	0.110	1.768	0.123	1.538	0.110
DHEP	8.793	0.568	16.199	1.124	27.114	1.945

shows the values of the Horwitz ratio in the ABIN samples fortified at different concentrations. Nonylphenols were excluded from this analysis, as they were not detected in the specified concentration range. Since the HorRat parameter in ABIN samples is less than 2, it can be generally said that the developed method is accurate.

LODs and LOQs were calculated using IUPAC criteria as 3.3 × S_b/m and 10 × S_b/m, respectively, where S_b represents the standard deviation of the y-intercept, while m is the slope of the calibration curve. The LODs of the selected analytes were between 0.6385 and 3.6275 mg/kg in ABIN and between 0.2072 and 5.1288 mg/kg in CTSH, and the LOQs were between 2.1284 and 12.0917 mg/kg in ABIN and between 0.6909 and 17.0961 mg/kg in CTSH. At concentrations of 1 mg/kg in cephalothorax + shells and 1.25 mg/kg in abdomen+intestine, all compounds were detected, except for nonylphenols.

4. Conclusions

The most relevant findings of this study are as follows. The analysis of eleven EDCs prone to contaminating shrimp has been conducted. This study represents the first instance of simultaneously determining all these analytes within a shrimp sample, thereby marking the inaugural examination of different sample components to verify the presence of contaminants in each. Furthermore, a method for the extraction of these 11 EDCs using the MSPD methodology and detection with HPLC-DAD has been successfully validated, achieving average recoveries of almost 90%. This demonstrates the merits of a versatile extraction method applicable to this type of matrix, which is also characterized by its simplicity, cost effectiveness, low solvent consumption and use of a green solid support, representing a significant advancement in research in this field.

Future research will prioritize the investigation of more environmentally benign solvents, such as natural deep eutectic solvents (NADES), to minimize the toxicity of the compounds and safeguard both environmental and operator health. One of the primary challenges will be the design and synthesis of eutectic solvents with low viscosity, enabling their effective use as extraction solvents. Additionally, the miniaturization and automation of the extraction technique will be investigated, and these advancements could result in a reduction in the use of chemicals, a lower energy consumption, and improved waste management.