Extraction and Analytical Techniques for Pharmaceuticals and Personal Care Products in Sediments: A Critical Review Towards Environmental Sustainability

Abstract

Pharmaceuticals and personal care products (PPCPs) are among the most frequently detected emerging pollutants in aquatic sediments, raising increasing concerns due to their persistence, bioaccumulation potential, and ecological impact. As sediments act both as reservoirs and secondary sources of contamination, effective and environmentally responsible analytical methodologies are essential for accurate environmental monitoring and risk assessment. This review presents a critical evaluation of extraction-based workflows for PPCP determination in sediment matrices, covering literature published from 2015 to 2025. We systematically analyze each step of the analytical pipeline, including sample pre-treatment, extraction, clean-up, and instrumental analysis, while emphasizing how method selection and optimization affect recovery rates, sensitivity, and detection limits. Special attention is paid to the physicochemical characteristics of PPCPs that govern extraction behavior, as well as to the trade-offs between analytical efficiency and environmental sustainability, such as solvent type, energy demand, and method greenness. By consolidating current knowledge, this work aims to lay a theoretical foundation for researchers and practitioners in selecting suitable, robust, and sustainable analytical strategies for effective environmental protection.

1. Introduction

Emerging pollutants (EPs) have emerged as a growing concern due to their intrinsic properties, including stability, persistence, and toxicity, which may negatively affect human health, as well as aquatic and terrestrial ecosystems [1]. Around 970 EPs have been detected by the NORMAN Network research group over the last 10 years [2]. According to Geissen et al. [3], at least 700 substances classified into 20 categories have been identified in the European aquatic environment. A considerable number of EPs derive from hospitals, household discharges, industrial activities, pharmaceutical manufacturing, animal husbandry, and wastewater. For instance, pharmaceuticals and personal care products, such as prescription and over-the-counter medications, cosmetics, detergents, and hygiene products, are widely used and frequently discharged into aquatic environments [4].

EPs, also known as micropollutants or contaminants of rising concern, represent a vast array of substances of synthetic or natural origin that are newly identified in the environment and are not typically included in routine monitoring programs. However, these pollutants are able to contaminate the environment with largely unknown effects on ecosystems and human health. Most of these compounds are not newly introduced but have been released into environmental compartments for many years; their presence, however, has only recently begun to be systematically investigated. Some are well-known pollutants already included in regulatory frameworks such as the Stockholm Convention (2001) on Persistent Organic Pollutants (POPs), the U.S. Clean Water Act list of Toxic and Priority Pollutants, or the EU Priority Substances list in surface waters [5]. EPs consist of organic or inorganic substances including a broad range of chemicals, such as deodorizers, fragrances, industrial chemicals, natural hormones, steroids, personal care products, pharmaceuticals, current-use pesticides, surfactants, and industrial additives or solvents [6].

Emerging pollutants disrupt aquatic ecosystems, harm wildlife, and cause reproductive, developmental, and neurological issues in humans. As a matter of fact, Ofloxacin reduces algal growth and photosynthetic rate, while Oxytetracycline weakens snail immune systems [7]. Water contamination by endocrine-disrupting chemicals (EDCs), particularly pharmaceuticals and personal care products, poses serious health risks such as thyroid dysfunction, Alzheimer’s disease, cancer, and obesity [8]. Several PPCPs, including triclosan, parabens, and phthalates, have been associated with endocrine-disrupting and hepatotoxic effects in aquatic organisms and humans, highlighting the need for comprehensive strategies to monitor and mitigate these pollutants [9].

The environmental release of such pollutants depends largely on their usage patterns and modes of application [10]. Their transport can be attributed to diffuse sources located in areas with concentrated water bodies [11]. EPs may reach aquatic systems via watersheds and subsequently be distributed through various pathways including groundwater, surface water, and seawater, ultimately ending up in sediments. In addition to diffuse sources, point sources such as wastewater treatment plants also contribute to the release of EPs into rivers, especially considering that conventional treatment processes are often ineffective in removing them. Typically, emerging pollutants are detected in water matrices at low concentrations [12]. However, a considerable number of analytical approaches also emphasize the detection of these pollutants in sediment matrices.

The results of accurate identification and categorization of EPs with respect to the employed analytical techniques in aquatic sediments may facilitate ecological risk assessment tools for the protection of marine ecosystems and, in turn, human health [13]. The merits obtained from such results can be translated into customized mitigation measures targeting EPs and their congeners, as well as the development of environmental policies and regulations to control their release into aquatic systems.

To support the establishment of effective regulations and policies for the understanding and management of EPs, continuous monitoring of their distribution in aquatic environments is required. In addition, identifying their physicochemical properties, fate, and behavior enables proper assessment of environmental impact and health risk. This, in turn, provides a baseline for developing mitigation strategies against potential hazards [14].

In this structured framework, indexed EPs along with appropriately selected sampling and analytical methods that ensure acceptable limits of detection (LOD), may be considered [3]. This allows for timely control of cumulative risks arising from their presence. LOD values and the corresponding recoveries are key metrics within quality assurance and quality control (QA/QC) processes, aiming to assess the accuracy (proximity to the true value) and precision (reproducibility) of the obtained results.

Regarding the risk assessment and determination of the environmental fate of EPs, a number of parameters are commonly used to analyze the occurrence of chemicals in water, providing crucial information on the parent EP compounds and their respective transformation products (TPs). In that sense, water solubility indicates a TP’s potential to accumulate in aquatic environments, while the octanol/water partition coefficient ($log{K}_{\mathrm{ow}}$) and the bioconcentration factor (BCF) reflect its lipophilicity and likelihood to bioaccumulate in organisms. Two additional key parameters are overall persistence (Pov) and long-range transport potential (LRTP), the latter representing migration possibilities of a target chemical. The impact of transformation on physicochemical properties and environmental behavior is a well-investigated subject. Further research, however, might shed light on accurately assessing the potential hazard that TPs and micropollutants derived from EPs pose to the environment and human health [15].

The focus of our survey is the determination of emerging pollutants (EPs) specifically related to pharmaceuticals and personal care products (PPCPs) in sediments found in various aquatic environments. These substances include a wide range of medicinal and hygiene-related compounds, many of which are classified as endocrine-disrupting chemicals (EDCs) due to their ability to mimic or interfere with hormonal function, potentially leading to adverse effects such as cancer and metabolic disorders [16].

Previous reviews have mainly examined pharmaceuticals and personal care products in surface waters or wastewater, emphasizing environmental occurrence or removal efficiency rather than analytical workflows. Studies such as aus der Beek et al. [17], Katsikaros and Chrysikopoulos [18], Kayode-Afolayan et al. [19], Abd El-Fattah et al. [20], and Boahen et al. [21] provided valuable overviews of contamination patterns and impacts but offered limited insight into sediment-specific analytical challenges or sustainability assessment. On the contrary, the present review focuses exclusively on sediment matrices, systematically evaluating 54 studies published between 2015 and 2025, comparing conventional and eco-friendly extraction approaches, and emphasizing solvent use, energy demand, and environmental sustainability. Furthermore, it introduces a structured taxonomy of extraction and determination workflows and discusses their degree of methodological harmonization.

Sediments serve as a sink where emerging contaminants (EPs) can accumulate due to their moderate to high lipophilicity, as indicated by octanol–water partition coefficients ($log{K}_{\mathrm{ow}}$) when they are found within 1.96–7.67. This phenomenon, referred to as the “sink effect,” signifies that sediments tend to retain significantly higher concentrations of EPs compared to the surrounding water, making them a potential secondary source of contamination. A representative case regarding PPCP accumulation is provided by Peng et al. [22], who investigated the presence of triclocarban (TCC) and its dechlorinated derivatives such as triclosan (TCS), as well as parabens and ultraviolet absorbents (UVAs) in sediment samples. The accumulation of these compounds may pose risks to wildlife, particularly benthic and sediment-dwelling organisms, as they can bioaccumulate in tissues and be transferred through aquatic food webs.

In line with green analytical principles, the selection of sample preparation and determination techniques should aim to minimize cost, analysis time, and organic solvent consumption, while maintaining high efficiency and selectivity for the target analytes through reduced matrix interference.

Sediment matrices have been analyzed using various extraction and analytical methods. Predominant extraction techniques include Soxhlet Extraction (SE), Pressurized Liquid Extraction (PLE), ultrasonication, and Solid-Phase Extraction (SPE). Traditional methods like SE remain widely used for extracting a broad spectrum of pollutants, while Gas Chromatography–Mass Spectrometry (GC-MS) is regularly employed due to its high sensitivity. In recent years, techniques such as PLE have gained popularity for their efficiency, automation, and reduced solvent requirements. Advanced instrumental approaches like Liquid Chromatography–Mass Spectrometry (LC-MS) and Ultra-High-Performance Liquid Chromatography with Tandem Mass Spectrometry (UHPLC-MS/MS) are prevalent for accurately quantifying PPCPs in sediments. These refined methodologies enhance the detection and analysis of emerging contaminants in complex sediment matrices.

Building on these observations, this work examines the occurrence and analytical determination of pharmaceuticals and personal care products in aquatic sediments. Considering the pivotal role of sediments in the fate and transfer of emerging pollutants, acting both as a sink and as a potential source of contamination, it is crucial to understand their analytical behavior in order to assess their environmental impact. To that end, we review related works published over the last decade (2015–2025), aiming to reveal critical aspects and trends of the state of the art concerning pre-processing, extraction, and subsequent steps of the analytical pipeline. Our main contribution lies in a systematic and thorough study of the proposed methodologies, in an attempt to fill certain gaps in the respective literature. Particular emphasis is placed on the influence of physicochemical properties of the target analytes on the selection of suitable extraction and determination methods. Furthermore, we introduce a structured and holistic taxonomy of the methodologies considered herein for the determination of PPCPs in various sediment matrices. In this framework, our goal boils down to establishing a solid theoretical standpoint for the development and selection of novel, efficient, and sustainable analytical strategies.

The structure of our work is as follows. In Section 2, we briefly present the searching methodology used to identify the related works considered herein. Section 3 overviews the generic steps of the detection and determination process for PPCPs in sediment matrices. In Section 4, we provide an in-depth analysis of the employed analytical techniques and propose a taxonomy of methods based on the extraction component of the analytical pipeline, along with the reported recoveries and limits of detection (LOD). Finally, conclusions are drawn and future directions are discussed in Section 5.

2. Searching Methodology

This review integrates existing scientific works on the extraction and analysis of pharmaceuticals and personal care products (PPCPs) as a major class of environmentally important analytes in sediment matrices. Candidate publications were identified across major academic repositories (Elsevier, Springer, MDPI, PubMed, ResearchGate, and Academia) using Google Scholar as the primary search tool. Searches were performed using combinations of terms referring to the analyte group (“pharmaceuticals”, “personal care products”, “PPCPs”), the environmental matrix (“sediments”), and the extraction or sample-preparation technique (e.g., Soxhlet, solid-phase extraction (SPE), pressurized liquid extraction (PLE), QuEChERS, microwave-assisted extraction (MAE), and accelerated solvent extraction (ASE)).

Approximately 150 studies published between 2015 and 2025 were initially retrieved. From these, 54 studies were retained for detailed comparative assessment according to the following inclusion criteria: focus on aquatic-sediment matrices; provision of complete methodological information (sample pre-treatment, extraction and determination procedures, recovery, and LOD/LOQ); English-language, peer-reviewed publication. Reviews, conference abstracts, and papers dealing exclusively with water, sludge, or biota were excluded. The selected studies were then organized following the taxonomy framework adopted in this review, encompassing analyte category, pre-treatment protocol, extraction parameters, instrumental method, and validation indices (recoveries, LOD/LOQ).

Figure 1 presents the temporal distribution of the selected PPCP-related studies over the last decade, along with the extraction techniques employed. A clear predominance of ultrasonication-based protocols (Ultrasonication) can be observed, followed by pressurized liquid extraction (PLE), and, at a lower frequency, microwave-assisted extraction (MAE), QuEChERS, and other less common approaches. Fluctuations in the number of studies per year likely reflect both emerging analytical trends and shifting research priorities in the environmental monitoring of sediment matrices. For a full description of these techniques, their acronyms, and detailed comparative data, the reader is referred to Section 4.1, Table 1.

It should be noted that the apparent reduction in the number of studies after 2020, as illustrated in Figure 1, stems from the rigorous inclusion criteria applied in this review. Only studies providing complete methodological and validation information were retained, ensuring consistent analytical coverage across the workflow.

3. Generic Workflow Pipeline of Emerging Pollutant Determination

In this section, we analyze each fundamental step of the methodologies, technologies, and processes involved in the determination of pharmaceuticals and personal care products (PPCPs) in sediment matrices. Sample pre-processing techniques, which are employed to facilitate the subsequent extraction as well as instrumental analysis phases, comprise the standard first step. Pre-processing mechanisms are essential for sample preparation, allowing samples to be efficiently exploited by the selected extraction approach.

With respect to the next step, the appropriate selection of the extraction method that best suits the task at hand heavily affects the upcoming determination task, in terms of the reliability and accuracy of the obtained results. Extraction is usually followed by a clean-up & instrumental analysis step, aiming to minimize interference from matrix components that are regularly conveyed in extraction results.

Finally, an instrumental analysis stage takes place to retrieve the determination results, namely compound identification and concentration, along with quality assurance and control steps concerning obtained recovery rates, corresponding limits of detection and quantification. Figure 2 illustrates the generic pipeline of these core steps for EP determination in aquatic sediments, some of which are examined in this work.

In essence, the sediment sample is immersed in the employed solvent (e.g., dichloromethane), and the extraction process is repeated for a standard amount of time to ensure adequate recovery. The prepared extract is then introduced into a determination step, often employing gas chromatography (GC) or liquid chromatography (LC) to separate the individual components of the extract based on their physical and chemical properties.

The separated compounds are usually ionized and fragmented in a mass spectrometer (MS), producing unique mass spectra for each compound. These spectra are then compared to reference databases, typically under standard quality assurance and quality control (QA/QC) conditions, in order to identify the detected compounds. Additionally, the intensity of the mass spectral peaks is used to quantify the concentration of the compounds in the sediment sample.

In the final step, average recoveries are calculated from spiked sediment samples, within defined limits of detection (LOD), to assess the overall efficiency and robustness of the analytical method.

3.1. Pre-Processing

This section highlights the procedures and techniques commonly employed for the storage, handling, and preservation of sediment samples as an initialization step for analyzing pharmaceuticals and personal care products (PPCPs) in aquatic matrices.

In the studies reviewed herein, sample preparation constitutes a critical initial stage of the analytical workflow. Intermediate steps before identification and quantification typically involve sediment preservation and pre-treatment through freeze-drying, lyophilization, or conventional air-drying to stabilize samples and prevent analyte loss.

Sample preparation comes in various forms depending on the nature of the sample, often requiring the majority of the total time spent during the analytical procedure. In several cases, this step significantly affects the quality of the obtained results and plays a critical role in the variability and discrepancies observed among otherwise similar approaches.

For instance, Álvarez-Ruiz et al. [23] suggest a series of pre-treatment steps in which samples are lyophilized, sieved, and homogenized prior to the extraction and clean-up stages. These procedures are part of a broader sediment manipulation process performed before chemical or toxicity testing. Sieving is conventionally applied to remove large particles and debris, whereas homogenization allows a larger sample to be evenly distributed and consistently used across multiple analytical tests.

Another factor that needs to be taken into consideration with regard to the storage and preservation of aquatic sediments is the initial conditions under which the sediments are sampled, transferred, and processed. Even slight disturbance during these stages may alter the bioavailability of the target contaminants.

Contamination may arise from container leaching, analyte sorption onto the container surface, oxidation, or degradation due to microbial activity. In addition, inappropriate container materials may also influence the accuracy of the retrieved analytical results.

Depending on the sediment type, its composition, and the employed analytical method, recommended storage standards usually involve dark conditions at temperatures below 4 °C [24]. Extended storage may lead to the loss of labile or volatile compounds (e.g., ammonia), or alter the sediment’s redox characteristics due to shifts in microbial activity.

Improper sample preservation can significantly affect analyte stability, leading to erroneous quantification and compromised data reliability. Therefore, the choice of storage conditions must ensure minimal chemical degradation and prevent volatilization, adsorption, or microbial transformation of target compounds.

At very low concentration, even the surface with which a sample comes into contact may act as a source of contamination. Such issues are frequently conveyed in the subsequent stages of the analytical process. Hence, careful sample collection and handling are needed to minimize external contamination sources and ensure analytical accuracy [25].

There are several standard techniques to ensure proper sample handling, including early refrigeration at controlled temperatures, the selection of suitable containers, and the addition of chemical preservatives. Although the complete elimination of contamination sources is unlikely, their impact can be minimized by following standardized procedures.

In [26], the authors denote the importance of choosing an effective sterilization method for laboratory experiments involving environmental samples, particularly in distinguishing between biotic and abiotic processes. The goal is to achieve sterilization that inhibits biological activity while preserving the physicochemical properties of the sample. The study compares three commonly used sterilization methods such as autoclaving, gamma irradiation, and sodium azide (${\mathrm{NaN}}_3$) and evaluates their effect on microbial abundance and activity in marine sediments. The same preservative is also employed by Cabrol et al. [27] to effectively suppress microbial activity in marine sediment samples.

Recent studies have highlighted that particle-size distribution and drying protocols significantly influence the physicochemical behavior of sediment samples and, consequently, their extraction performance. Ouendi et al. [28] demonstrated that fine fractions (<0.063 mm) possess the highest adsorption capacity and organic-matter content, while oven-drying leads to partial loss of reactive components compared with air- or freeze-drying. Zentar et al. [29] further showed that oven-drying alters the Methylene Blue Value (MBV), Atterberg limits, and organic-matter levels, producing coarser apparent grain-size distributions and reduced adsorption potential; freeze-drying was therefore recommended for preserving sediment integrity and representativeness. Similarly, Wu et al. [30] reported that different drying treatments (freeze, air, oven) induce mineralogical transformations of amorphous Fe and Al oxides into more crystalline forms, reducing adsorption capacity and increasing bioavailable phosphorus fractions. Collectively, these findings emphasize that fine, organic-rich fractions govern sediment reactivity and that standardized freeze- or air-drying protocols are essential to maintain natural composition and ensure reproducible extraction efficiency.

In a nutshell, pre-processing steps such as drying, grinding, and homogenization are relatively standard procedures, consistently applied in a similar manner across related studies. Limited attention has been devoted to their further improvement, partly due to the inherent difficulty of optimizing these steps. As a result, the main focus of the research community has shifted towards the selection and development of more efficient extraction techniques.

3.2. Extraction

This section examines the most widely used extraction techniques proposed for the determination of emerging pollutants (EPs) in aquatic sediments, with a primary focus on pharmaceuticals and personal care products (PPCPs). Current research efforts focus both on optimizing existing methods and developing new extraction strategies, aiming to minimize matrix interferences and eliminate co-extracted compounds that could compromise analytical performance. When properly selected and optimized, extraction methods can provide the accuracy and sensitivity required for reliable quantification of target analytes.

Efficient extraction is typically associated with satisfactory recovery rates and low detection limits, enabling trace-level analysis. Several studies have reported high performance in terms of accuracy and sensitivity in the determination of target compounds from environmental solid matrices using various extraction protocols [31,32,33,34,35,36]. In the following, extraction methods are broadly classified into solvent-based and sorbent-based categories, as applied to aquatic sediments.

Solvent-based methods utilize organic solvents to extract analytes from sediments, based on their solubility to isolate target compounds. These techniques comprise a wide range of protocols such as Liquid–Liquid Extraction (LLE), Microwave-Assisted Liquid–Liquid Extraction (MLLE), QuEChERS (Quick, Easy, Cheap, Effective, Rugged, and Safe), Microwave-Assisted Extraction (MAE), Ultrasonic-Assisted Extraction (UAE) or Ultrasonication (UT) or Ultrasonic Extraction (USE)(in what follows we adopt UAE as the standard terminology for this extraction method), Pressurized Liquid Extraction (PLE), also known as Accelerated Solvent Extraction (ASE) (for consistency we keep PLE in what follows), Soxhlet and Soxtec extraction, Solidified Floating Organic Drop Microextraction (SWE-DLLME), Thermal Desorption (TD), Stir Bar Assisted Extraction (SAE), Supercritical Fluid Extraction (SFE),Sequential Extraction Procedure (SEP), and Direct Solvent Extraction (DSE).

Each method offers specific advantages depending on the physicochemical properties of the analytes, the complexity of the matrix, the required extraction efficiency, and the instrumental analysis that follows. The selection of an appropriate solvent-based extraction technique is thus guided by multiple performance and sustainability criteria, including solvent volume, energy demand, and compatibility with downstream detection methods.

While solvent-based methods typically provide exhaustive extraction and high recoveries, they often require larger solvent volumes, longer processing times, and more extensive clean-up. In contrast, sorbent-based approaches generally yield cleaner extracts with reduced matrix interferences, though their performance depends strongly on sorbent selectivity, conditioning, and compatibility with the target analytes.

On the other hand, sorbent-based methods rely on solid-phase materials that selectively retain target analytes while removing interfering matrix components. This category includes techniques such as Solid-Phase Extraction (SPE), Solid-Phase Microextraction (SPME), and Stir Bar Sorptive Extraction (SBSE), which are widely employed for both sample preparation and clean-up [37].

These methods typically involve passing the sample through a sorbent medium (e.g., cartridge or disk), where the analytes are selectively adsorbed. The matrix is subsequently removed through washing, and the retained compounds are eluted with an appropriate solvent. Additional techniques within this category include Solvent Precipitation-Liquid Extraction (SPLE), dispersive Solid-Phase Extraction (dSPE), Molecularly Imprinted Solid-Phase Extraction (MISPE), and Matrix Solid-Phase Dispersion (MSPD).

These additional techniques differ not only in operational principles but also in their environmental footprint and potential for automation, underscoring the need to balance analytical efficiency, sustainability, and throughput when designing extraction workflows.

In what follows, we review the most commonly applied extraction methods for emerging pollutants (EPs) in aquatic sediments with a particular focus on PPCPs, highlighting the underlying principles of each technique and, where possible, discussing their respective advantages and limitations.

3.2.1. Distinction Owing to Optimization Parameters

As noted in Section 3.1, sample preparation depends on the nature of the target analytes, the characteristics of the matrix, and the separation technique employed. Since no universal sample preparation protocol is suitable for all sample types [38], quantitative analysis in complex matrices usually requires a targeted strategy to address the wide array of co-existing contaminants. These factors can hinder the analytical process or interfere with the signal response of the target compounds.

Conventional extraction techniques, such as liquid–liquid extraction, Soxhlet, Soxtec, UAE, and mechanical shaking, have long been employed for the recovery of emerging pollutants from solid or liquid environmental matrices. However, these methods are increasingly regarded as obsolete due to their intensive use of organic solvents, large sample and reagent consumption, and significant waste generation. Their time-consuming protocols and environmental burden have led to a gradual shift towards more sustainable alternatives.

To address these issues, a range of novel, greener solvent extraction methods have emerged over the last two decades. These include Solid-Phase Extraction (SPE), Solid-Phase Microextraction (SPME), Pressurized Liquid Extraction (PLE), Supercritical Fluid Extraction (SFE), Matrix Solid-Phase Dispersion (MSPD), and Microwave-Assisted Extraction (MAE), all of which aim to reduce solvent usage, processing time, and environmental impact.

Sample preparation remains a critical step in the development of effective extraction protocols, especially when target analyte concentrations are low. In this context, recent trends in solvent extraction follow two key directions: (i) the search for environmentally friendly solvents, and (ii) the miniaturization of extraction systems.

An illustrative example of the first trend is Pressurized Hot Water Extraction (PHWE), a PLE-based method using water as a green solvent. In PHWE, water undergoes physicochemical changes under high temperature and pressure, enhancing its extraction efficiency as a highly polar solvent [39]. This technique is often combined with Stir Bar Sorptive Extraction (SBSE) to reduce contamination risks. Similarly, PHWE coupled with SPE and ultra-performance liquid chromatography–tandem mass spectrometry (UPLC-MS/MS) has been successfully applied for multi-residue analysis in solid samples [40].

Another promising solvent family comprises ionic liquids (ILs), which consist of organic cations and organic or inorganic anions. ILs can dissolve a wide range of substances and are characterized by low volatility and reduced toxicity, making them attractive alternatives to conventional organic solvents [41].

A third alternative involves the use of surfactant-based extraction media, such as non-ionic surfactants in Cloud-Point Extraction (CPE) and ionic surfactants in Coacervative Extraction (CAE). These systems are associated with lower toxicity, minimal volatility, and, in some cases, enhanced biodegradability [42].

Miniaturization refers to the scaling down of extraction protocols often in combination with energy-assisted techniques such as UAE, MAE, SFE, and PLE. These methods facilitate the transfer of analytes into aqueous or organic phases and are widely applied for the extraction of organic compounds from solid samples. In addition to enhancing extraction efficiency, they offer substantial benefits in terms of reducing solvent volumes, waste generation, processing time, and operational cost. Current developments focus on automation and further miniaturization, frequently involving solvent-free or low-solvent systems that pre-concentrate analytes directly in the acceptor phase [43].

These methodologies are particularly suitable for the extraction of trace-level pollutants, such as PPCPs, from solid environmental matrices. They combine high extraction efficiencies with improved sustainability and reduced environmental footprint. Over the last two decades, the scientific community has devoted increasing attention to minimizing the toxic and hazardous nature of conventional organic solvents. As a result, analytical strategies have increasingly emphasized the use of green solvents and the adoption of miniaturized, eco-friendly extraction workflows.

Comparing extraction techniques for pharmaceuticals and personal care products (PPCPs) in solid matrices is inherently challenging due to the heterogeneity and binding characteristics of these samples. Solid and semi-solid sediments often require exhaustive extraction to release analytes tightly associated with the matrix. Method selection therefore depends primarily on analyte properties, matrix composition, and solvent efficiency, with solvent type and extraction duration being key optimization parameters.

Three operational modes define the way the extraction procedure for solid matrices can be carried out when targeting emerging pollutants (EPs), including PPCPs:

i.

Static mode: A fixed volume of extractant is used. This mode normally requires simpler instrumentation and lower solvent consumption [44].

ii.

Dynamic mode: The extractant follows a continuous flow through the sample [45], allowing improved mass transfer, faster solvent movement through the matrix, and reduced compound degradation, especially in challenging extraction situations.

iii.

Hybrid mode: Combines the merits of static and dynamic modes [46], aiming to achieve high efficiency while maintaining practical extraction times.

Most extraction techniques, such as LLE, LSE, SPME, and PLE, operate in the static extraction mode and are widely applied in the analysis of emerging pollutants (EPs), including PPCPs. Apart from static mode, methods such as MASE, SAE, and MAE can also support dynamic operation. Among other parameters, such as temperature, pressure, and extraction time, the choice between dynamic and static extraction mode can also influence the selectivity and extraction efficiency of PHWE methods. PHWE can be performed in either mode [47]. In the static mode, solvent consumption is limited to a fixed volume. By contrast, in dynamic extraction, a larger volume of fluid is required to reach equilibrium, as fresh solvent is continuously pumped through the sample.

A common disadvantage of a dynamic system, compared to its static counterpart, lies in its inherent complexity. In some cases, extraction efficiency can be lower, since the contact time between the solvent and each unit of sample is short. Nevertheless, dynamic extraction offers notable benefits. The continuous flow of fresh solvent into the sample improves extraction kinetics, and a filtration unit can be readily installed in the system, eliminating the need for separate filtration and rinsing steps after extraction. Moreover, a clean-up module can often be integrated to further reduce total extraction times. Dynamic extraction is particularly suitable for labile analytes, including certain classes of emerging pollutants (EPs) such as PPCPs, since they are flushed directly from the system and thus degradation caused by high temperature or pressure is minimized [48].

Apart from the standard distinction of extraction techniques into solvent-based and sorbent-based, Albaseer et al. [25] also highlight the use of supercritical fluid extraction (SFE), which constitutes a distinct category within gas or fluid-based extraction techniques. SFE [49] is often considered an environmentally friendly approach, as ${\mathrm{CO}}_2$ is used as the extraction medium. This eliminates the need for large volumes of organic solvents, which are otherwise often required for the efficient extraction of (semi)polar pharmaceuticals and personal care products (PPCPs). Another advantage of SFE is that clean-up can be readily incorporated into the extraction step. The main drawback, however, is that optimizing the extraction conditions can be challenging.

Several solvent-based methods, such as Soxhlet extraction, are still widely used for the analysis of pharmaceuticals and personal care products in solid matrices. Soxhlet remains an attractive option for routine analysis due to its robustness and relatively low cost. However, the extracts obtained are usually unclean and require extensive, time-consuming clean-up. For this reason, Soxhlet is more suitable when only a few samples are processed, and when results are not required on the same day of analysis. Overall, it belongs to a family of highly labor-intensive and time-demanding techniques.

Although green solvent systems (e.g., PHWE, ILs, DES) and miniaturized extraction approaches (e.g., UAE, MAE, PLE) share the common objective of reducing environmental impact, they differ considerably in analytical robustness and field applicability. Green solvents often provide superior environmental profiles and compatibility with polar PPCPs, but their extraction kinetics and long-term stability can be difficult to control. Conversely, energy-assisted methods achieve faster and more exhaustive extraction with high reproducibility, yet may require additional clean-up and instrument-specific optimization. Therefore, the most sustainable analytical workflows will likely arise from hybrid strategies that combine low-toxicity solvents with miniaturized, automated extraction systems offering both eco-efficiency and analytical reliability.

3.2.2. Key Modern Extraction Techniques for Solid Environmental Matrices

In the following, we present an overview of key modern extraction techniques applied to solid environmental matrices, with applicability to a wide range of emerging pollutants (EPs), including PPCPs. The aim is to pick out the factors and operational parameters that contribute to high extraction efficiency, while also considering solvent consumption and sustainability aspects.

Pressurized solvent extraction (PSE) is a solid–liquid extraction technique proposed as an alternative to conventional approaches such as Soxhlet extraction, maceration, percolation, or reflux. It offers notable merits in terms of reduced solvent consumption and improved extraction efficiency [50]. Pressurized–liquid extraction (PLE), has proven to be a rapid and efficient method for extracting PPCPs from a wide variety of solid samples [51]. Additionally, microwave-assisted extraction (MAE) and sonication-assisted extraction (SAE) have been widely applied to solid environmental matrices [52,53]. Both PLE and MAE are considered among the most exhaustive extraction techniques available. They often achieve high recoveries, even for PPCP analytes that are tightly bound to the matrix.

Particularly, PLE is well suited for solid environmental samples containing PPCPs, especially when the target analytes are tightly bound to the matrix. Such efficient extraction, however, often yields highly contaminated extracts that require further purification. One way to tackle this drawback is to combine sample clean-up with PLE by adding suitable adsorbents directly into the extraction cell.

The extraction efficiency of PLE is strongly influenced by pressure and temperature settings, as well as by matrix effects. As a consequence, its extraction behavior can be complex, and optimizing the operating conditions is often labor-intensive. Another limitation is that, when hydrophobic organic solvents are used, high water content in the sample significantly decreases analyte recovery, as water obstructs contact between the solvent and the analyte [54].

In [55], PLE was applied in an automated configuration, achieving short extraction times (approximately 15 min), low solvent consumption (15–40 mL), and eliminating the need for post-extraction filtration. The main drawbacks reported were the high equipment cost and the substantial optimization effort required to avoid efficiency bias owing to matrix dependency.

Microwave-assisted extraction (MAE) involves placing the sample with the extraction solvent in specifically designed containers and heating the solvent using microwave energy. This technique is particularly appropriate for routine analysis of PPCPs, wherein time and solvent consumption are considerably reduced, while high sample throughput can be achieved. MAE has been successfully applied for the extraction of organic contaminants, including PPCPs, from solid and semi-solid environmental matrices.

From a moderate equipment cost perspective, MAE is considered feasible, as it is generally less expensive than contemporary extraction methods such as SFE. In addition, MAE involves only minor safety risks, since most extractions are carried out under atmospheric conditions. However, there are certain drawbacks. Non-polar solvents, which are poor absorbents for microwave heating [56], are discouraged in MAE applications. The technique also exhibits relatively low selectivity, often attributed to its dependence on solvent nature and extraction temperature, which is not always desirable for PPCPs. Furthermore, MAE may lead to the loss of active PPCP compounds during additional, time-consuming clean-up steps, which do not always preserve the full range of extracted analytes [57].

Nevertheless, MAE techniques are typically highly efficient. The simultaneous transfer of heat and mass in MAE creates a cooperative effect that accelerates extraction and improves overall yield. In the context of pharmaceuticals and personal care products, MAE is also regarded as a green technology due to its reduced consumption of organic solvents [58].

When comparing the volume of organic solvents required, both PLE and MAE outperform traditional extraction approaches, with the latter generally consuming less solvent. Regarding total analysis time, an increasingly important factor, MAE is also faster than PLE. Although MAE requires additional cooling and filtration before obtaining the final extract, it allows concurrent extractions in a microwave oven, whereas PLE can only perform sequential runs. Moreover, prior to extraction, PLE demands more labor-intensive and time-consuming sample preparation, as the dispersion of the sample with diatomaceous earth requires weighing the material and homogenizing the mixture with a mortar [54].

Under optimized conditions, Pressurized Liquid Extraction (PLE) and Microwave-Assisted Extraction (MAE) generally achieve similar or higher recoveries than conventional techniques such as Soxhlet for sediment samples containing PPCPs.

UAE is based on the application of high-frequency sound waves and a limited amount of solvent to efficiently extract target compounds. Within the ultrasound range, two main regions can be distinguished: (a) power ultrasound (20–100 kHz), characterized by high intensity and commonly used for extraction and processing applications, and (b) signal or diagnostic ultrasound (100 kHz–10 MHz), employed primarily in clinical diagnostics as well as in quality control and assessment [59]. In the context of PPCPs, UAE offers the potential for effective recovery of analytes while using reduced solvent volumes.

The improvement in extraction yield during UAE is primarily attributed to the formation of cavitation bubbles, which are generated and compressed during sonication. The rapid compression increases the temperature (approximately 4500 °C) and pressure (about 50 MPa), causing the bubbles to collapse. This process enhances solvent penetration into cells and increases the surface area of contact between the solid and liquid phases [60]. In PPCP applications, these effects facilitate the efficient release of analytes that may be tightly bound within complex matrices.

The extraction efficiency in UAE is influenced by multiple parameters, including temperature and extraction time, solvent polarity, the amount of extractant, sample mass and type, and ultrasound frequency and intensity, as well as the number of pulses applied. In recent years, the technique has gained considerable attention for PPCP analysis due to its notable advantages over conventional approaches: low solvent volumes, short extraction times, low instrumental cost, and minimal economic and environmental impacts. Its application under laboratory conditions is straightforward, while its feasibility at industrial scale is also promising, owing to the reduced duration of the procedure, increased extraction yields and selectivity. Moreover, it allows enhanced mass and heat transfer.

Despite the successful use of ultrasound in extraction, several drawbacks of ultrasonic cavitation have been reported. These include the potential degradation of thermolabile PPCP compounds due to high temperatures and ultrasound intensity [61]. The literature also notes that the mode of action can be complex, with efficiency closely linked to the structural properties of the solid matrix and the composition of the liquid phase. In some cases, degradation of target analytes by free radicals may occur. Moreover, the method assumes that the extraction solvent must efficiently absorb acoustic energy, which is not always guaranteed [62]. Additional limitations include the need for filtration steps and the generation of high noise levels during operation.

Matrix solid-phase dispersion (MSPD) is an alternative sample preparation technique, particularly suited for solid and semi-solid samples [63]. It is a manual method that combines sample disruption with the dispersal of the sample onto particles of a solid sorbent. The merits of MSPD include simplicity, low cost, short extraction time, and reduced organic solvent consumption. In addition, it operates under mild conditions (atmospheric pressure and room temperature), making it flexible and convenient while enabling simultaneous extraction and clean-up. These features have extended the application of MSPD to various classes of compounds, including PPCPs, pesticides, pollutants, and other analytes from complex samples such as animal tissue, plant material, and environmental matrices [64].

Nonetheless, MSPD can be relatively tedious, as the sample must be ground with the solid matrix and then packed into a column for extraction. This requirement can be a deterrent for many applications, particularly when large volumes of solvent are still needed for extraction and clean-up.

While the fundamental principles of on-line and off-line sample preparation are shared across different application fields, their practical adoption is strongly matrix-dependent, with aqueous systems offering greater compatibility with on-line integration than complex solid matrices such as sediments.

A promising direction in modern analytical strategies for PPCPs involves the automation and integration of sample pre-treatment within chromatographic systems, particularly for aqueous matrices, where on-line coupling between extraction/clean-up and the separation–detection stage is technically feasible and has been widely adopted. On the contrary, for complex solid matrices such as sediments, sample preparation remains predominantly an off-line process. In such workflows, exhaustive extraction and clean-up are performed prior to chromatographic analysis, often involving solvent-based extraction followed by solid-phase extraction (SPE) or other fractionation steps.

This reliance on off-line protocols in sediment analysis is exemplified by recent studies, such as that by Leite et al. [65], who validated a UHPLC–TOF–MS method for pharmaceuticals in coastal sediments following conventional off-line extraction and clean-up. Although on-line configurations have been demonstrated for certain solid matrices other than sediments (e.g., SPME–GC, SFE–LC, SFE–GC, PHWE–LC, PHWE–GC, SAE–LC, SAE–GC, MAE–GC) [38], their application to PPCP analysis in sediments remains extremely limited, primarily due to the need for exhaustive extraction and matrix clean-up prior to analysis, which yields complex extracts that are difficult to transfer directly into an on-line workflow.

In general, on-line sample preparation integrates extraction and clean-up directly with the chromatographic system, allowing the entire extract to be transferred without intermediate handling. When performed in a closed, automated configuration, it can improve sensitivity and reduce the risks of contamination, analyte loss, and degradation. The trade-offs include higher system complexity and lower flexibility in method adaptation.

Solid-phase extraction, whether implemented in an off-line or on-line configuration, involves partitioning the analytes between a solid sorbent and a liquid phase, with the analytes exhibiting greater affinity for the sorbent than for the sample matrix. Coupling SPE directly to LC systems in an on-line mode has gained considerable attention for PPCP analysis in aqueous samples [66]. For sediment matrices, however, SPE is generally performed as an off-line clean-up following exhaustive extraction, sometimes in combination with microporous membrane liquid extraction (MMLE) [67,68].

Irrespective of whether on-line or off-line strategies are adopted, extraction performance for PPCPs is affected by numerous factors, including but not limited to the choice of extraction materials (adsorbents), operating conditions, solvents, agitation methods (e.g., micro-extraction), and the separation and detection technologies employed. Looking ahead, future research is expected to focus on developing low-cost extraction methods that offer simple operation, minimal equipment requirements, and functional extraction materials capable of enhancing analyte recovery and selectivity.

Overall, the comparative assessment of extraction approaches for PPCPs in sediments indicates that no single technique universally outperforms others. Instead, method applicability depends on a compromise between matrix characteristics, target compound properties, and analytical objectives. Traditional exhaustive solvent-based extractions offer robustness and reproducibility, whereas miniaturized and green alternatives provide sustainability and rapid throughput at the cost of increased optimization effort. A critical synthesis of the literature suggests that hybrid or integrated workflows which combine mild extraction conditions with selective sorbents or in-cell clean-up modules, represent the most promising direction toward high-efficiency and environmentally benign analytical protocols.

3.3. Clean-Up & Preconcentration

After extraction, a clean-up step usually takes place to remove matrix components that can interfere with the subsequent chromatographic or mass spectrometric analysis. Such interferences may include lipids, pigments, sterols, or, in some cases, elemental sulfur. Common post-extraction clean-up methods involve adsorption on alumina, florisil, or silica gel columns. For sulfur-rich marine sediments, copper is frequently employed to remove sulfur, as these matrices often originate from reducing environments [69]. Although the general principles are similar across analyte classes, clean-up protocols for PPCPs must be carefully tailored to avoid loss of polar or labile compounds during the removal of co-extracted matrix constituents, as recoveries for certain PPCPs after SPE clean-up can vary markedly depending on sorbent type and elution conditions [70].

Clean-up methods mainly rely on liquid–liquid extraction (LLE) or fractionation using column chromatography or solid-phase extraction (SPE). These are considered classical techniques and remain widely used for the preconcentration and clean-up of environmental samples. Among existing approaches, LLE and column chromatography are particularly tedious, as they typically involve multiple manual steps and require large volumes of organic solvents. Consequently, these procedures usually include an additional concentration step by solvent evaporation. In the following, we highlight the most widely employed clean-up techniques applied in the analysis of sediment samples.

Regarding SPE, high costs and time-consuming procedures are expected. On the contrary, LLE is considerably less expensive [71] whereas the solvent extract needs to be dried by evaporation [72].

SPME can also serve as a clean-up method in sediment analysis. Relying on a fiber coated with a specific sorbent material, it selectively extracts target analytes while minimizing matrix interferences. This simple and efficient sample preparation technique has gained significant popularity across diverse application fields. It can be deemed a green approach, as it minimizes extensive sample handling and reduces solvent consumption. In addition, SPME can be readily integrated with mass spectrometry (MS), enabling rapid and potentially real-time analysis of the extracted analytes. This integration not only allows for short determination times but also accommodates automation of the entire analytical process [73].

Dispersive solid-phase extraction (dSPE) has also gained considerable utilization, offering notable advantages such as streamlining the clean-up process, enabling the simultaneous analysis of multiple samples, providing rapid results, and reducing solvent consumption. In dSPE, a solid sorbent—typically composed of silica or polymer—is added directly to the sample solution, combining the extraction and purification steps into a single stage [74].

Solid-phase extraction with dual molecularly imprinted polymers (SPE-DMIP) is a widely used technique for the purification of heterogeneous sample types, including sediments. A potential drawback of SPE-DMIP is the additional time and effort required during method development and optimization, compared with conventional SPE approaches. Nonetheless, SPE-DMIP offers enhanced extraction efficiency, selectivity, and ease of use, rendering it a valuable tool in sediment analysis.

Centrifugation can be employed to separate solid particles present in sediments from the liquid phase, thereby enabling the removal of undesirable particulate matter prior to analysis.

Filtration is a straightforward and frequently used approach for eliminating larger particles from sediment samples. Filters with different pore sizes can be selected according to the desired degree of particle removal.

Gel permeation chromatography (GPC) is a size-exclusion chromatography technique that separates compounds based on their molecular size. It is particularly useful for removing high-molecular-weight or interfering substances from sediment extracts. Straightforward automation and reduced risk of damage to analytical instruments are among its pros. However, it requires large volumes of solvents whereas the extracts may still contain co-extractives from the matrix. Another con is related to the fact that the use of solvents in GPC can generate significant amounts of hazardous waste [75].

Column chromatography methods use adsorbents such as Florisil or Silica Gel to separate and remove interfering compounds from sediment extracts. By these means, the compounds of interest are eluted separately from the column. Column chromatography methods employ adsorbents such as Florisil or Silica Gel to separate and remove interfering compounds from sediment extracts. In this way, the compounds of interest are eluted separately from the column.

The clean-up step plays a critical role in removing matrix interferences such as lipids, pigments, and sulfur compounds that can distort chromatographic responses. Brennan et al. [76] compared gel permeation chromatography (GPC) and solid-phase extraction (SPE) for sediment extracts containing fipronil and its metabolites, reporting comparable recoveries (72–119%) and detection limits (0.12–0.52 µg ${\mathrm{kg}}^{-1}$ d.w.). Although GPC efficiently removed high-molecular-weight compounds, optimized SPE with mixed sorbents (PSA/GCB, Florisil/GCB) achieved similar performance with lower cost and solvent demand. Massei et al. [77] integrated a sequential clean-up step using silica and alumina directly within pressurized liquid extraction (PLE), effectively minimizing ion-source contamination during LC–HRMS analysis of multi-polar compounds. Muijs et al. [78] demonstrated that inadequate purification leads to inflated baseline signals due to residual organic matter; their comparative evaluation of sorbents confirmed that partially deactivated Florisil (5 %) and alumina (10 %) yielded the most consistent recoveries (∼95%) while eliminating lipid interferences. In summary, these studies denote that optimized clean-up, whether implemented as a separate or integrated step, is indispensable for accurate quantification, particularly in organic-rich sediment matrices.

It should be noted that extraction and clean-up may, at least to some extent, influence key validation parameters under investigation (e.g., recovery rate, limits of detection, limits of quantification). All in all, clean-up and pre-concentration are two essential steps in analytical sample preparation: the former removes unwanted interferences and contaminants, while the latter increases the concentration of target analytes. Both contribute directly to improving the accuracy and sensitivity of the analytical method.

3.4. Instrumental Analysis

The potency of separative techniques such as chromatography, especially when coupled with MS for the analysis of target compounds is considered as a fundamental aspect of contemporary instrumental analytical chemistry. In 1952, GC became the most employed among separative techniques owing to the seminal work of James and Martin on partition chromatography [79]. According to Snyder et al. [80], it has been estimated that only about $20\%$ of organic molecules are GC-amenable, followed by a limited number of molecules which can be volatilized under tardy, high-cost, and often, irreproducible derivatization tasks. LC can deal with the remaining part of the molecules for separation, without being constrained by volatility or thermal stability.

In an attempt to compare GC and LC, the desorption method of analytes from the stationary phase is the main difference between them. Reliable and fast results are an appealing feature of GC methods. Although being dynamic in its development phase, GC involves shortcomings such as long analysis times and challenging conditions for real-time or direct quantitative analysis. However, gas chromatography–mass spectrometry (GC–MS) is a significant analytical technology widely used across many applications and fields. Typically, GC–MS analysis takes 20–40 min per sample [81], which is much longer than common LC methods to date. Lehotay et al. [82] reported that LPGC–MS is the most practical and advantageous fast-GC technique, as it can achieve analyses in under 10 min.

The efficiency of LC–MS/MS systems can be improved through various optimization strategies, including the use of smaller-particle-size columns, increasing LC flow rates, utilizing specialized stationary phases, and employing mass spectrometric techniques such as selected Multiple Reaction Monitoring (MRM). Another widely used optimization approach is multiplexing, in which target analytes from multiple LC streams are introduced into a single mass spectrometer. This technique can substantially increase the throughput and overall analytical capacity of the LC–MS/MS system [83].

The appropriate selection of an analytical method for a given sample is strongly influenced by the choice of extraction solvent. Different solvents exhibit varying affinities for specific analytes, depending on their physicochemical properties. For example, some analytes may display higher solubility in certain solvents than in others. A solvent with high selectivity for the target analytes will generally yield greater extraction efficiency and sensitivity for the selected analytical method. Black and co-workers [84] evaluated a novel extraction approach, Energized Dispersive Guided Extraction (EDGE^®), for the analysis of 210 pesticides in complex solid matrices. They compared the efficiency of two extraction solvents, i.e., acetonitrile and hexane:acetone (1:1, v/v), and applied graphitized carbon clean-up with three different elution solvents: Acetonitrile, dichloromethane (DCM), and DCM:ethyl acetate (EtOAc) (1:1, v/v).

Additionally, different analytical methods require specific sample preparation procedures. For instance, GC is better suited for volatile and thermally stable compounds, while LC favors more polar and non-volatile compounds. The choice of extraction solvent should align with the characteristics of the analytical method to ensure optimal performance and accurate results. Thus, it is crucial to select an extraction solvent compatible with the sample matrix to achieve efficient analyte recovery. The type of matrix (e.g., water, soil, sediment, biological tissues) also affects extraction efficiency since some solvents dissolve target analytes more effectively in a given matrix, whereas others may be less suitable.

Having studied the physicochemical properties of analytes and understood the behavior of different extraction solvents, researchers can make informed decisions when selecting the most suitable analytical method for a given sample and analyte of interest. Integrating knowledge of both the analyte and the extraction solvent can improve the sensitivity, accuracy, and reliability of analytical measurements.

Our review of analytical methods proposed over the last decade indicates that the majority of approaches for aquatic sediments involve liquid chromatography–tandem mass spectrometry (LC–MS/MS), UHPLC–MS, HPLC–DAD–ESI–MS, UPLC–MS/MS, and UHPLC–Q-Orbitrap HRMS, and, very rarely, GC–MS/MS. The appropriate choice of analytical method depends on the nature of the target analyte, including emerging contaminants, with particular emphasis on PPCPs.

3.5. Quality Assurance and Quality Control (QA/QC)

During the experimental procedure for identifying and quantifying analytes of interest, it is essential to ensure that specific requirements related to the reliability of the results are met. Such requirements include quantifying extraction efficiency based on surrogate standards, or assessing the accuracy of recovered spiked analytes using blank matrix spikes. These measures are applied throughout the analytical procedure to ensure that results fall within acceptable limits. In other words, a quality management system is responsible for ensuring that these requirements are satisfied to achieve the desired quality of results.

Within this framework, quality assurance (QA) and quality control (QC) are interrelated components of the quality management system, typically implemented through procedural blanks, spiked blanks, spiked matrices, sample duplicates, surrogate standards, and other measures. QA aims to ensure that the required quality will be achieved, whereas QC verifies that the desired quality has been met. QA focuses on preventing unreliable or biased results, while QC seeks to detect and identify potential sources of bias.

In practice, QA encompasses all methodologies used to ensure that laboratory results are reliable. Analytical quality control (AQC) processes are particularly important for environmental analyses, especially when analyte concentrations are extremely low and close to the detection limit. In laboratories with well-defined procedures, AQC processes are often embedded into routine operations, for example through the random introduction of known standards into the sample stream or the use of spiked samples. Ultimately, AQC comprises all tasks that ensure results are consistent, comparable, and accurate, falling within specified limits of precision. QC follows the entire analytical workflow, from sample collection to final data reporting.

All in all, QA is the verification system that ensures the entire analytical testing process remains within acceptable limits. QC comprises the processes established to achieve accurate results efficiently, while also quantifying any unreliability when results fall outside acceptable limits. Figure 3 summarizes the main components of a quality management system, outlining the requirements to be met at each step of the analytical pipeline.

QA is supported by internal quality control (IQC) and external quality control (EQC) programs. IQC focuses on individual analytical methods and assesses their performance against mathematically derived quality criteria. Standard methods often include extensive validation data and are frequently approved by credible international or national organizations. Before implementing a method for routine use, it must be properly validated, at minimum by assessing:

i

Linearity: Define the calibration range and demonstrate a linear response if possible. If calibrants do not exhibit a linear response, appropriate data transformation should be considered.

ii

Limit of detection (LOD): Determine the lowest analyte concentration distinguishable from zero with 95% confidence.

iii

Precision: Evaluate within-day and between-day coefficients of variation at three concentration levels.

iv

Accuracy: Where possible, analyze certified reference materials or perform inter-laboratory comparisons.

EQC assesses the accuracy of analytical methods by comparing results from one laboratory with those from others analyzing the same material. This is typically achieved by a reference laboratory distributing specimens with known and unknown concentrations to participating laboratories, which analyze the samples and report results back. EQC reports should clearly indicate whether performance is satisfactory. If not, the cause must be identified and corrected, and the IQC program reviewed to address any shortcomings. The overarching goal of EQC is to improve inter-laboratory comparability and provide independent verification of accuracy. Participation in EQC programs is recommended for all routinely analyzed variables, provided IQC is also an integral part of the laboratory’s procedures.

Recent studies have emphasized the importance of transparent QA/QC reporting and its direct impact on the accuracy and comparability of analytical data. Lescord et al. [85] highlighted that inconsistent or incomplete QA/QC documentation, such as missing blanks, unreported recoveries, and lack of calibration validation, remains a major limitation in environmental analyses and can obscure inter-laboratory comparability, leading to under- or over-estimation of pollutant levels. In PPCP extraction workflows, practical examples further illustrate how method-specific QA/QC directly affects analytical reliability. Sadutto et al. [86] demonstrated that variations in sorbent chemistry and pH control during solid-phase extraction (SPE) led to analyte-specific recovery losses and strong matrix effects, revealing the need for standardized conditioning procedures. Darwano and co-workers [87] reported that inadequate compensation for matrix interferences during ultrasound-assisted extraction resulted in poor recoveries (<60%) for compounds such as diclofenac and sulfamethoxazole, until corrected through a standard-addition calibration. By contrast, Langford et al. [88] exemplified effective QA/QC implementation by maintaining procedural blanks below detection limits, thereby confirming that reported PPCP concentrations reflected genuine environmental levels. Collectively, these studies indicate that insufficient or poorly documented QA/QC measures can distort analyte recoveries, elevate apparent detection limits, and compromise data comparability. On the contrary, transparent and systematic QA/QC validation which covers recovery, precision, and matrix-effect correction, remains fundamental for reproducible and defensible sediment analyses. In this framework, thorough QA/QC validation across all workflow stages forms the foundation upon which effective quality-assurance frameworks operate.

Overall, QA achieves these objectives by establishing protocols and quality criteria for all aspects of laboratory work, while providing a framework within which IQC and EQC programs can operate effectively. In essence, QA is primarily a management system and is analyte-independent, as it concerns the overall operation of the laboratory rather than individual analytical tasks.

4. Taxonomy of PPCPs in Sediments

In this section, we review the literature on pharmaceuticals and personal care products (PPCPs) detected in aquatic sediments. Each study is categorized according to the extraction component of the analytical pipeline, highlighting how methodological choices and intermediate analytical processing steps influence recovery, sensitivity, and overall analytical performance.

PPCPs comprise a diverse group of chemical compounds intended for personal health or cosmetic purposes, as well as agriculture-oriented products designed to enhance livestock growth or well-being [89,90]. Continuous research and monitoring are of paramount importance to evaluate potential risks to human and ecosystem health, as pharmaceuticals, together with their metabolites and degradation products, are consistently introduced into the environment.

In recent decades, studies worldwide have shown that various PPCPs accumulate in aquatic sediments originated from, e.g., marine [16], lagoon [91], lake [92], river [31] or dam [66] environments. Müller et al. [93] studied the potential routes of exposure to endocrine disrupting chemicals (EDCs) for fish living in sediments under field conditions.

Conducting studies to assess PPCPs in various environmental compartments is crucial for determining their distribution, identifying contamination sources and establishing limit values to safeguard the environment and human health. To that end, a careful analysis of their physicochemical parameters is essential, since they provide valuable information on their behavior, fate and transport in the environment. Some of the physicochemical parameters which are important in the analysis of these contaminants include solubility, vapor pressure, octanol–water partition coefficient ($K_{ow}$), melting point, boiling point, Henry’s law constant, and pH. As a matter of fact, PPCPs with higher solubility in water may have a higher likelihood of being dissolved in the water phase and subsequently transported to sediments. On the other hand, compounds with lower solubility tend to adsorb onto sediment particles, leading to their accumulation in sediments.

Traditional methodologies such as Solid–Liquid Extraction (SLE), Solid-Phase Extraction (SPE), Soxhlet Extraction, and Supercritical Fluid Extraction (SFE) have been widely used in the past. Due to their limitations in terms of time, cost, and environmental impact the research interest is shifted towards simplification, speed, cost-effectiveness, and reduced use of hazardous solvents to render them environmentally friendly compared to conventional techniques. Concerning PPCPs our study, which is summarized in Table 1, concludes that most of the traditional approaches employ Soxhlet which allows for the extraction of a broad range of pharmaceuticals, including both active pharmaceutical ingredients and their metabolites, from sediment matrices.

In modern techniques however, more refined approaches have gained popularity such as Pressurized Liquid Extraction (PLE). PLE is an automated and efficient extraction technique that utilizes elevated temperatures and pressures to enhance the extraction process. It is particularly useful for extracting pharmaceuticals from sediment samples due to its ability to efficiently extract a wide range of analytes with different chemical properties. It allows for the extraction of both polar and non-polar compounds, which are commonly found in pharmaceuticals. The automation and versatility of PLE make it a preferred choice in modern pharmaceutical analysis, as it offers increased efficiency, reduced solvent consumption, and improved extraction performance compared to traditional methods such as Soxhlet Extraction or SPE.

With respect to the analytical methods used for the determination of PPCPs, our work considers LC-MS and UHPLC-MS/MS to be the most prevalent due to the intrinsic physicochemical parameters of the PPCPs. These techniques rely on the separation and identification of target compounds based on their physicochemical properties, allowing for the quantification of pharmaceuticals in sediment samples.

Before discussing specific extraction families, it is useful to outline the geographical and methodological scope of the reviewed literature. Figure 4 provides an integrated overview of regional activity and downstream analytical practices across the 54 studies analyzed. Nearly half of the reviewed works originate from Europe (40.7%) and more than one third from Asia (35.2%), with smaller but noteworthy contributions from North America (9.3%) and Africa (9.3%). This regional distribution reflects both the solid analytical infrastructure and the more established monitoring frameworks in European and Asian laboratories.

Regarding sample cleanup, solid-phase extraction (SPE) dominates the literature (50.0%), followed by direct post-extraction analysis without additional purification (40.7%). The latter category, labeled as “None”, corresponds to workflows in which high-selectivity MS/MS or HRMS instrumentation mitigates matrix interferences without a distinct cleanup step. Minor occurrences include dSPE and hybrid sorbent combinations such as PSA or on-line SPE (<5% each). Instrumentally, LC–MS/MS platforms prevail (37.0%), complemented by UHPLC–MS/MS (13.0%) and a variety of advanced high-resolution or hybrid MS systems (e.g., QqLIT, TOF, Orbitrap, QqQ) collectively representing about 50% of the remaining studies. This distribution confirms that tandem and high-resolution mass spectrometry constitute the analytical backbone of PPCP determination in sediments.

In the following, we exhaustively discuss the analytical steps of the proposed works examined herein so as to determine the efficiency of each distinct extraction method according to the observed recovery rates at defined limits of detection.

4.1. Family of Approaches in Emerging PPCPs

In this Section, we present a taxonomy of the different approaches proposed over the last nine years with respect to the employed extraction methodology as the primary categorization component, followed by determination and recovery stages of the analytical pipeline for various types of emerging PPCPs in sediments.

To facilitate interpretation, Figure 5 summarizes the hierarchical logic adopted for this taxonomy. The reviewed studies are first classified according to the employed extraction method (e.g., PLE, MAE, UAE, SPE, Soxhlet, QuEChERS, etc.), which serves as the primary key of the analysis. A secondary grouping follows based on the analytical determination technique (e.g., GC–MS, LC–MS/MS, UHPLC–ESI–MS, HPLC–DAD) and performance indicators such as recovery and LOD/LOQ values. The figure also integrates key sediment pre-treatment steps and physicochemical attributes of PPCPs that govern method selection. Finally, it provides a light guidance zone suggesting the general suitability domains of extraction approaches according to PPCP polarity and sediment composition. This schematic visually connects the classification used in Table 1 with the methodological discussion presented in Section 3.2, offering a concise overview of how extraction, determination, and validation stages interact throughout the analytical pipeline.

In this context, we group distinct categories of approaches in Table 1 over a wide variety of PPCPs’ classes, such as antibiotics [31,37,94,95,96,97,98,99,100,101,102,103,104,105,106,107], illicit/abused drugs [23,32,108], non-steroidal anti-inflammatory drugs (NSAIDs) [91,100,103,107,109,110], anti-epileptic drugs [104,109,111], endocrine disrupting compounds (EDCs) [16,112,113,114], anti-bacterial drugs [107,115], $\beta$-blockers [91,100,116], veterinary medicine (anthelmintic) drugs [111,117], cardiovascular drugs [103,118], and other overlapping groups of PPCPs [4,86,101,119,120,121].

PLE is a sample preparation technique that uses solvents under elevated temperature and pressure to extract target analytes from solid or semi-solid samples. Accelerated solvent extraction (ASE) is a commercial implementation of PLE, and the two terms are often used interchangeably. Thirteen antibiotics, including azithromycin (AZI), erythromycin (ERY) and clarithromycin (CLA) [31], 19 antibiotics (ABs) [94], 20 illicit drugs [32], 16 PPCPs [107], a class of anti-bacterial (triclocarban) [115], 36 out of 56 PPCPs (antibiotics, analgesics, anti-arrhythmic, anti-epilepsy, anti-lipidemic, anti-tumor, blood vessel dilators, and others) [96], and a number of chiral drugs comprising anti-depressants, $\beta$-blockers, $\beta$-agonists, anti-histamines, and stimulants (fluoxetine, amphetamine, propranolol, venlafaxine, and citalopram) [116] were extracted using PLE, followed by LC coupled with tandem MS in most cases. In [107,115], for example, coupling was performed with HRMS.

Antibiotics such as azithromycin (AZI), erythromycin (ERY) and clarithromycin (CLA) are regularly prescribed for the treatment of chest infections (e.g., pneumonia), dermatological conditions (e.g., cellulitis), and other bacterial infections including otitis media and Helicobacter pylori-associated ulcers. During method development, both ultrasonic extraction (UAE) and pressurized liquid extraction (PLE) were systematically evaluated; however, the optimized and validated protocol employed PLE with $0.2\%$ ${\mathrm{NH}}_3$ in MeOH, whereas USE was applied solely during the preliminary solvent screening phase for antibiotics in river sediments [31]. An analytical methodology based on PLE extraction and SPE clean-up, followed by LC–MS/MS determination has been validated and applied to assess the occurrence of 20 abused drugs and their metabolites in 144 sediment samples collected in four Spanish river basins [32]. Therein, satisfactory recoveries between $90\%$ and $135\%$ were achieved at $1.1$ ng/g (d.w.) LOD. Triclocarban is an anti-bacterial active ingredient widely used for personal cleaning products including deodorant soaps, deodorants, detergents, cleansing lotions, and wipes. The use of triclocarban restricts the transmission of germs to other people or to objects. They are usually detected in wastewater effluents and they can accumulate in sediments. Depending on the circumstances, the overall method recoveries were $89\%$ at $0.01$ to $0.1$ ng/g LOD [115]. Recent advancements reported in [107] extend these methodologies to extract 16 PPCPs, including antibiotics (ciprofloxacin, enrofloxacin, amoxicillin, erythromycin, sulfamethoxazole), anticonvulsants (carbamazepine), NSAIDs (diclofenac, ibuprofen, aspirin), analgesics/antipyretics (paracetamol), stimulants (caffeine), antibacterial and antifungal agents (triclosan), insect repellents (N,N-diethyl-meta-toluamide, DEET), and hormones (17$\beta$-estradiol, 17$\alpha$-ethynyl estradiol), from freeze-dried surface marine sediment using PLE in a solvent mixture of methanol:ultrapure water (1:1, v/v) at 80 °C for three cycles. Obtained recoveries ranged from $60\%$ to $100\%$, with LOD of $0.1$–$0.3$ ng/g and LOQ of $0.3$–$0.9$ ng/g. More recently, Teysseire et al. [122] developed a method for the analysis of 13 pharmaceuticals and 5 personal care products in freeze-dried lake sediment samples. Extraction was performed using PLE at 80 °C, followed by SPE for clean-up. Two successive extractions were carried out using methanol and a methanol:water mixture (1:2, v/v). Analysis was conducted using liquid chromatography coupled to a triple quadrupole mass spectrometer (LC-QqQ-MS), yielding recoveries ranging from $12.6\%$ to $108.1\%$, with LODs between $0.01$ and $3.75$ ng/g and LOQs from $0.04$ to $12.51$ ng/g.

36 PPCPs were extracted using PLE in a solvent extraction of $\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v) in pH = 11, a step prior to LC-MS/MS to obtain a good recovery of 85.7–124.6% [96]. Pharmaceutically active compounds (PhACs), including carbamazepine (an anticonvulsant and mood stabilizer), ciprofloxacin, and sulfamethoxazole (both antibiotics), were analyzed in lagoon sediments following lyophilization. Extraction was performed using PLE, and clean-up was carried out with Oasis HLB SPE cartridges. Quantification by LC–MS/MS yielded detection and quantification limits of $0.25$ and 1 ng/g for carbamazepine, and $0.5$ and 2 ng/g for ciprofloxacin, respectively [123]. An analytical methodology based on ASE-Chiral LC–MS was successfully developed to determine pharmaceutical and chiral drugs. In this case, sufficient recoveries up to $22\pm 3\%$ for R(-)-chlorpheniramine and $93\pm 5\%$ for acebutolol-E1 were achieved [116].

It should be noted that the molecular weight and polarity of PPCPs can affect their mobility and persistence in sediments. Actually, heavier compounds with higher molecular weight might have a lower tendency to migrate within the sediment matrix compared to lighter compounds. In a similar fashion, the polarity of PPCPs can influence their affinity for sediment particles with more polar compounds having a higher probability of sorption. For this reason, it is important to understand the physicochemical properties of pharmaceuticals in order to select the appropriate extraction method so as to gain efficient recoveries. For instance, the octanol–water partition coefficient ($K_{ow}$), can provide information about the tendency of PPCPs to partition between different phases. Hence, compounds with higher $K_{ow}$ values tend to preferentially accumulate in sediment organic matter, while those with lower $K_{ow}$ values may exhibit higher mobility in the aqueous phase.

To highlight the significance of monitoring the physicochemical properties, we refer the reader to the study of Chabilan et al. [94] which showed that the $log{K}_{\mathrm{ow}}$ values of 19 antibiotics (ABs) are important in the process of their extraction. Therein, PLE was employed, followed by solid-phase extraction and liquid chromatography and subsequently coupled with tandem mass spectrometry, from river sediments. As a matter of fact, the study suggested to order ABs based on their $log{K}_{\mathrm{ow}}$ values so as to facilitate the selection of the most suitable extraction solvent. $K_{ow}$ with $log{K}_{\mathrm{ow}}$ values below zero yielded lower recoveries than positive $log{K}_{\mathrm{ow}}$ values whereas each $log{K}_{\mathrm{ow}}$ group required different extraction conditions (e.g., solvent, pH). The method as assessed at a relative recovery of the analytes and surrogates within $45\%$ and $125\%$.

Microwave-assisted extraction (MAE) is another extraction technique which utilizes microwave energy to generate heat within the sample, promoting the release and extraction of target compounds, as opposed to the use of elevated temperature and pressure of PLE. Their differences can be summarized in the following factors:

i

Heating method: While PLE uses externally applied heat in a pressurized environment, MAE relies on microwave irradiation to generate heat in the sample.

ii

Temperature and pressure: Accurate control of both temperature and pressure are feasible in PLE, offering more flexibility in optimizing extraction conditions. In MAE, the temperature is typically controlled by the microwave power and exposure time.

iii

Solvent: In MAE, solvents are typically added to the sample to enhance extraction efficiency. Instead, the use of pressurized solvents that are circulated through the sample matrix, facilitate the extraction of target compounds in PLE.

iv

Automation/scalability: PLE systems are typically automated and can process multiple samples concurrently, offering higher throughput. MAE configurations can also be automated but are often used for smaller-scale extractions.

Despite those differences, both MAE and PLE offer advantages such as faster extraction times, improved efficiency compared to conventional extraction methods. The choice between MAE and PLE, as well as other modern approaches, depends on the specific analytical requirements, target compounds, sample matrix, and available equipment.

Among the various pharmaceuticals regarded as emerging pollutants, benzimidazoles, a sub-class of anthelmintic drugs, such as flubendazole and fenbendazole, are of particular concern because of their large-scale use in veterinary medicine and their health effects on aquatic organisms. For this reason, it is essential to have reliable analytical methods which can be used to simultaneously monitor their appearance and determine their concentration in environmental matrices, such as river sediments. Recently, Rojewska et al. [124] developed MAE–SPE–LC–MS/MS methods validated using marine sediment samples. Extraction and determination involved two steps: MAE under controlled conditions (400 W, 60 °C, 10 min), followed by solid-phase extraction (SPE) for clean-up. Two extraction methods were tested: Method 1, based on a mixture of 10% magnesium nitrate, saturated ammonium chloride, and methanol; and Method 2, employing methanol:dichloromethane (4:1, v/v) under acidic pH. Determination was carried out via LC–MS/MS, and extraction performance was assessed through absolute recovery (AR). Recovery values varied depending on the extraction method and sediment characteristics, ranging from below 20% to approximately 80%, with Method 2 generally offering improved recovery for analytes affected by matrix-related ion suppression. Wagil et al. [117] have employed MAE for the determination of flubendazole (FLU) and fenbendazole (FEN) using a mixture of hexane:acetone (1:1, v/v), followed by liquid chromatography–tandem mass spectrometry (LC–MS/MS) analysis. Reported absolute recoveries of flubendazole and fenbendazole were between $98.3\%$ and $103.4\%$.

UAE refers to the use of ultrasound waves to enhance the extraction of compounds from solid or liquid samples. Ultrasound is applied to the sample, leading to the formation and collapse of small bubbles through a process called cavitation. This phenomenon generates localized high temperatures and pressures, facilitating the release of target compounds from the sample matrix and improving extraction efficiency.

Wu et al. [125] extract 21 pharmaceuticals from river sediments using sonication, centrifugation and SPE clean-up. Sulfamethoxazole (SMX) showed stronger sorption onto sediments compared with other antibiotics due to its higher pseudo-partitioning coefficients (846–10,786 L/kg). The extracted compounds are analyzed using LC–MS/MS. A mixed solution of NaCl (sodium chloride), commonly used as a salting-out agent, enhances the extraction of certain compounds by promoting the partitioning of analytes into the liquid phase. Oxalic acid is a weak organic acid that can act as a chelating agent, helping to dissolve and extract metal ions and other polar compounds from the sediment. Lastly, ethanol is used so as to dissolve numerous of organic compounds. The combined effect of the solvents and sonication enhances the extraction efficiency, thus allowing for the release of the target analytes from the sediment and their subsequent partitioning into the solvent.

In [126], sonication is employed to extract 6 pharmaceuticals from sediment samples, followed by SPE to clean-up and concentrate the extracted pharmaceuticals before the LC-MS/MS analysis. Reported recoveries and LOD ranged from $40.1\%$ to $120.6\%$ and from $0.02$ to $2.5$ ng/g, respectively.

USE was employed for the detection of twenty-two antibiotics in river sediment samples. Acetonitrile and citric acid buffer (pH 3) were used as extraction solvents, followed by LC-MS/MS analysis [95] yielding 53–149% recoveries. The same extraction and analytical approach were employed in [108] for 11 abused drugs in freeze-dried river sediments, wherein 10 mL of McIlvain buffer–methanol (1:1, v/v) was used to achieve matrix spike recoveries between 68–101% at LOD between 0.20–1.50 ng/g. Similarly, the work of Monteiro and colleagues [66] was based on an LC-MS/MS system for the determination and quantification of 12 antimicrobials in lyophilization dam sediments. Ultrasonication was again proposed as an effective extraction technique along with on-line SPE clean-up to obtain successful recoveries from $89\%$ to $119\%$ at LOD between 0.40–5.1 $\mathsf{\mu }$g/kg.

Reliable methods were also developed for the simultaneous extraction and determination of 32 antibiotics [97] and ARVDs [98] in sediments using similar extraction protocols employing a McIlvaine buffer with ACN (1:1, v/v) and $\mathrm{Mg}{\left({\mathrm{NO}}_3\right)}_2$-${\mathrm{NH}}_3·{\mathrm{H}}_2\mathrm{O}$ (96:4, v/v), as well as MeOH and ${\mathrm{H}}_2\mathrm{O}$ as extraction solvents, respectively. Specifically, Chen and co-workers performed a comprehensive optimization of SPE clean-up parameters, i.e., the appropriate selection of SPE cartridge (GCB + HLB, SAX + HLB), dilution volumes (where 350 mL proved sufficiently efficient on average) and the addition of ${\mathrm{Na}}_2\mathrm{EDTA}$ (wherein 0.2 g ${\mathrm{Na}}_2\mathrm{EDTA}$ were statistically significant in improving the recoveries of antibiotics). In this respect, recoveries of the target compounds ranged between 40–$127\%$, and $84.3$–$111.3\%$, respectively. A multi-residue methodology based on Ultrasound-assisted extraction, SPE clean-up and LC–MS/MS analysis was developed for the determination of highly abused illicit drugs and some of their metabolites in particulate matter, sewage sludge and sediments. 21 of the 41 target compounds are investigated in these environmental solid matrices [23]. In [127], Nantaba et al. reported the analysis of 25 pharmaceutical compounds in lake sediments, including 14 antibiotics (trimethoprim, sulfamethoxazole, sulfamethazine, sulfacetamide, oxytetracycline, tetracycline, erythromycin, roxithromycin, ciprofloxacin, levofloxacin, norfloxacin, sparfloxacin, enoxacin, and metronidazole), four anti-epileptic and antidepressant drugs (carbamazepine, fluoxetine, diazepam, and salbutamol), three analgesic and anti-inflammatory agents (ibuprofen, diclofenac, and acetaminophen), three beta-blockers (propranolol, atenolol, and metoprolol), and one lipid regulator (bezafibrate). Extraction was performed using UAE with acetonitrile and pH 3 citric acid buffer for 15 minutes, followed by SPE clean-up. Quantification was carried out using LC–MS/MS, achieving recoveries of 70–$111\%$, with LODs ranging from $0.3$ to $3.2$ ng/kg and LOQs from $1.2$ to $11.2$ ng/kg.

In [109], a UE–LC–PDA method has been successfully developed and applied for the analysis of pharmaceutical compounds in soil and sediments. SPE was also employed as an additional clean-up step prior to the analysis. Reported recoveries ranged from $74\%$ to $112\%$ and LOD from $0.010$ to $0.027$ $\mathsf{\mu }$g/kg. In another study, 10 PPCPs were extracted from river sediments, including 2 stimulants (paraxanthine, caffeine), 2 antibiotics (sulfamethoxazole, acetyl sulfamethoxazole), 2 antiepileptics (carbamazepine, epoxide carbamazepine), and 4 NSAIDs (ibuprofen, carboxyibuprofen, diclofenac, 4-hydroxy diclofenac). Extraction was performed using UAE with two cycles of acetone and two cycles of methanol for 25 min. Ethyl acetate showed good recovery for acidic compounds but poor performance for basic ones. Acetone improved overall recovery but was still limited for basic analytes, while methanol enhanced recovery of basic compounds but reduced that of acidic ones. The acetone–methanol combination was found to be optimal, achieving recoveries of 69–$109\%$, with LODs in the range $0.848$–$2.756$ $\mathsf{\mu }$g/kg and LOQs in the range $2.828$–$9.192$ $\mathsf{\mu }$g/kg, using liquid chromatography–photodiode array detection (LC–PDA) for analysis [128].

QuEChERS (Quick, Easy, Cheap, Effective, Rugged, and Safe) is a sample preparation method that combines several steps into a single procedure, including sample homogenization, liquid–liquid extraction, and dispersive solid-phase extraction. A salt induces phase separation whereas sorbents remove unwanted matrix components, thereby achieving analyte enrichment and clean-up at the same time. QuEChERS is favorably known for its simplicity, speed, and cost-effectiveness compared to traditional extraction methods. It has gained attraction in the field of food analysis, particularly for multi-residue analysis of pesticides, due to its efficiency and robustness.

85 pharmaceuticals and personal care products (PPCPs) in sediments were analyzed in [129], wherein the sediment was spiked with internal and surrogate standards. QuEChERS was employed along with 10 mL of acetonitrile and 10 mL of purified water (Milli-Q water, or MQW) as extraction solvents. Additionally, 6 g of anhydrous ${\mathrm{MgSO}}_4$, which acts as a drying agent, and 1.5 g of sodium acetate, a pH adjuster, were also added to the mixture. This agitation facilitated the extraction of the PPCPs from the sediment into the solvent. 1.2 g of ${\mathrm{MgSO}}_4$ and 0.4 g of primary–secondary amine (PSA) served as a clean-up procedure in the QuEChERS method. As a secondary clean-up step, the acetonitrile phase containing the extracted compounds was transferred to 15 mL tubes for dispersive solid-phase extraction (dSPE). The recovery rates for the surrogate compounds in the sediment samples were determined. The recovery rates were found to be $67\pm 9\%$ for sulfamethoxazole-13C6, $108\pm 11\%$ for alprazolam-d5, and $104\pm 17\%$ for benzophenone-d10. These percentages indicate the effectiveness of the extraction and analysis methods in successfully recovering these surrogate compounds from the sediment samples.

QuEChERS and simultaneous dispersive solid-phase extraction (dSPE) clean-up were proposed for blood lipid regulators, analgesics, anti-inflammatory drugs and $\beta$-blockers in lagoon sediment samples [91]. The extracts were clean enough for determination by (LC-MS/MS). In this case, recoveries lied between $27\%$ and $120\%$ and LOD ranged from $0.001$ to $0.75\mathrm{ng}/\mathrm{mL}$. A simple, rapid, and rigorous method using a modified QuEChERS approach followed by dSPE clean-up and LCLIT/Orbitrap MS analysis was proposed, allowing the simultaneous determination of 25 PPCPs in sediments. In light of these considerations, satisfactory recoveries between $64\%$ and $101\%$ were achieved [119].

Although the research interest is generally shifted towards modern approaches that favor speed, cost-effectiveness, environmentally friendly techniques, etc., traditional extraction methods sometimes cannot be neglected, especially in cases where high recovery rates are expected. Actually, there are several modern approaches that derive from modifications applied on traditional approaches.

For instance, Ismail et al. [16] and Cha et al. [37] employ traditional extraction methods, i.e., Soxhlet and SPE clean-up, as well as, SPE extraction, respectively, with liquid chromatography–tandem mass spectrometry (LC-MS/MS). In the former approach (EDCs), average recoveries range within 50.39–129.10%, whereas the latter reports generally higher recovery (>75%) of 4 BLs and 4 PEs from all samples, except for amoxicillin (AMOX).

Traditional solid-phase extraction (SPE) typically involves the use of solid sorbents, such as silica or polymer-based materials, to selectively extract and purify target analytes from a sample matrix. A modified version of SPE, dubbed dispersive solid-phase extraction (dSPE) is considered as a modern extraction method. Conventional SPE typically involves the use of solid sorbents, such as silica or polymer-based materials, to selectively extract and purify target analytes from a sample matrix. dSPE on the other hand, is a newer approach that simplifies the traditional SPE procedure by using a dispersive step wherein the sorbent material is mixed directly with the sample extract or solution containing the analytes of interest. The sorbent particles adsorb the target analytes, while unwanted matrix components are left in the solution. After mixing and adsorption, the sorbent is separated from the solution, either by centrifugation or filtration, and the analytes are eluted from the sorbent for subsequent analysis. dSPE offers advantages such as simplicity, speed, and reduced solvent consumption compared to traditional SPE methods. Moreover, it is usually embedded in other extraction methodologies. For instance, QuEChERS includes a dSPE step. It has gained popularity, particularly in the field of sample preparation for analysis of contaminants in food and environmental samples.

An extraction method was successfully optimized and validated for the analysis of 40 antibiotics from cephalosporin, fluoroquinolone, lincosamide, macrolide, nitroimidazole, quinolone, sulfonamide and tetracycline groups [99]. A mixture of MeOH and MeCN (1:3 v/v) extraction solvents was used in dSPE, prior to LC-MS/MS analysis. In this case, satisfactory recoveries between 24–162%, 48–151%, 51–159%, and 50–149% were achieved for spiking levels 10, 20, 50, and 100 $\mathsf{\mu }$g/kg, respectively.

Apart from the widely used LC-MS/MS approach for detecting pharmaceuticals in sediments, there is also a growing interest in Ultra-High-Performance Liquid Chromatography with Tandem Mass Spectrometry (UHPLC-MS/MS) due to its high sensitivity, selectivity, and capability to analyze complex samples.

An analytical method combining pressurized liquid extraction (PLE) and solid-phase extraction (SPE) as pre-treatment process coupled with UHPLC-MS/MS was developed for the simultaneous determination of 19 anthelmintic drugs in environmental sediment samples [111]. 34 pharmaceuticals have been detected by ultrasonicated extraction and centrifugation with SPE clean-up, followed by UHPLC-MS/MS analysis. Satisfactory recoveries between $43\%$ to $118\%$ were achieved as well as LOD from $0.01$ to $0.6$ ng/g [4]. In a similar extraction methodology fashion and analysis: (i) 7 Pharmaceuticals, 1 Personal Care Product and 3 Hormones in dried river sediments, (ii) 4 different groups of pharmaceuticals: Anti-inflammatory (Ketorolac, Naproxen), antibiotics (Ofloxacin, Ciprofloxacin), anti-cancer (Ifosfamide, Cyclophosphamide), $\beta$-Blockers (Atenolol, Propranolol) and (iii) 111 organic micropollutants (OMPs), including antiepileptic, antineoplastic agents and antihistamine, in air-dried lake sediments were determined in [92,100,130], respectively, with satisfactory recoveries between 54.0–94.4%, 87–113%, and 81–104%, respectively. The method in [130] demonstrated several key qualities, including suitable linearity, low detection and quantification limits, good repeatability and inter-day precision, absence of ion interference, and negligible carryover and matrix effects. Ultrasonic extraction, optimized through sequential solvent use and centrifugation, achieved high recoveries (>60% for most compounds). The method’s limit of detection ($0.66$ ng/g for all pharmaceuticals) was lower than values reported in other studies for similar, less complex matrices such as soil, sewage sludge, broiler manure, agricultural soil and sediments.

UAE and SPE were used to pre-treat 25 types of Pharmaceutically active compounds (PhACs) in 6 categories which were detected in marine sediments: 9 antibiotics, 9 hormones, 3 NSAIDs, 2 antipsychotic drugs, 1 hypoglycemic drug and 1 antiviral drug. The method combined ultra-high-performance liquid chromatography and electrospray ionization-tandem mass spectrometry (UHPLC-ESI-MS/MS) for the detection of target compounds. In this work, satisfactory recoveries ranging from $61\%$ to $117\%$ were achieved [101].

A multiresidue analytical method for the determination of 22 endocrine-disrupting compounds (ECDs) in sediments has been proposed in [112]. The UAE procedure followed by SPE with graphitized carbon black (GCB) cartridge as the clean-up step were used. The final extract was analyzed by ultra-high-performance liquid chromatography coupled with tandem mass spectrometry (UHPLC-MS). The use of GCB for SPE with single clean-up yielded excellent recoveries, ranging between $75\%$ and $110\%$. Pharmaceutical compounds in marine surface sediments were extracted using UAE with a MeOH:ACN (2:1, v/v) mixture for 15 min at 25°C, followed by clean-up with Oasis HLB SPE cartridges. Detection and quantification were performed using ultra-performance liquid chromatography coupled with quadrupole time-of-flight mass spectrometry (UPLC–QTOF-MS). The method achieved recoveries ranging from $75\%$ to $105\%$, with LODs between $0.3$ and $3.9$ ng/g, demonstrating high sensitivity and suitability for trace-level pharmaceutical analysis in marine sediments [131]. Additionally, 30 PPCPs in river sediments were analyzed using UAE with 10 mL of acetonitrile:deionized water (1:1), followed by SPE clean-up. Analysis by UPLC–Orbitrap/MS yielded recoveries of 74–$109\%$, with LODs from $0.02$ to $1.21$ ng/g and LOQs from $0.5$ to $14.5$ ng/g [132].

In another multiresidue method [102], QuEChERS and SPE clean-up, followed by UHPLC-MS/MS analysis, yielded good recoveries for most of the 27 psychiatric drugs and antibiotics. Psychiatric drugs had recoveries from $47.2\%$ (sertraline) to $104\%$ (diazepam) whereas sulfonamide and fluoroquinolone antibiotics had lower recoveries, below $30\%$ and $20\%$, respectively. The recovery results for antibiotics varied depending on the antibiotic family. In sediments, the use of basic acetonitrile as the extraction solvent resulted in good recoveries for psychiatric drugs and macrolide antibiotics. The uncharged form of these compounds had a higher affinity for the extraction solvent. Trimethoprim showed a recovery of $84.4\%$, while macrolides erythromycin, clarithromycin, and azithromycin had recoveries higher than $70\%$. However, sulfonamide and fluoroquinolone antibiotics exhibited lower recoveries since the strong interaction between these antibiotics and sediments made their extraction challenging.

Two solid-phase extraction methods were systematically studied to determine 32 pharmaceuticals and personal care products in sediments by UHPLC-MS/MS. One involves HLB cartridges activated with sodium dodecyl sulfate (SDS) before the passage of the sample to form an ion pair with cationic analytes, while the other uses mixed HLB–cation exchange cartridges. Under those circumstances, $60\%$ of the compounds had recoveries greater than $80\%$ and LOD equal to 10 ng/g [86].

In sediment analysis, solid–liquid extraction (SLE) refers to the transfer of target analytes from a solid matrix into a suitable liquid phase through direct contact with an extracting solvent. This approach is versatile, allowing the use of a wide range of solvent systems and operational modes, and can be readily coupled with clean-up steps such as solid-phase extraction (SPE) prior to instrumental analysis. Depending on the solvent composition and extraction conditions, SLE can be suited to improve recoveries for both polar and non-polar compounds.

A solid–liquid extraction (SLE) which uses a mixture of methanol and water (1:1, v/v) solution was proposed in [118]. The supernatant was diluted with water and cleaned up by stir-disc solid-phase extraction (SPE). A specially designed buckypaper disc made of carbon nanotubes is used as the sorbent membrane for extraction. The disc is activated with nitric acid to enhance its extraction capability for polar compounds. Again, all extracts were analyzed by UHPLC–MS/MS. The method showed good recovery rates for the analytes, with ∼$69\%$ recovery for low spike levels and ∼$80\%$ recovery for medium and high spike levels. Only a few analytes exhibit recovery values below $50\%$. The relative standard deviation (RSD) of the method is consistently below $20\%$, indicating good precision. Overall, reported recovery ranged within 50–$111\%$ and LOD between $0.02$–$9.9$ ng/g. In [65], Leite et al. present the development and validation of a sensitive UHPLC–TOF–MS method for detecting 30 pharmaceuticals in marine sediments, in accordance with Commission Regulation (EU) 2021/808. Solid–liquid extraction was performed using 4.5 mL of acetonitrile and 0.5 mL of water. The method achieved recoveries ranging from $74.3\%$ to $115.4\%$, with LODs between $0.02$–$0.23$ ng/g and LOQs from $0.06$–$0.69$ ng/g.

High-Performance Liquid Chromatography with Mass Spectrometry (HPLC-MS) is also often used as an alternative to UHPLC-MS/MS for the analysis of PPCPs in sediments. The selection of one technique against the other usually depends on the specific requirements of the study, the desired sensitivity, throughput, selectivity, identification needs, scope of analysis and available resources. PLE followed by HPLC–MS analysis has been proposed as a potent method to determine 46 pharmaceutical products (PPs), in marine sediments [103] with reported recovery >$75\%$.

The same HPLC–MS analysis of pond sediments for extracting pharmaceuticals—naproxen (NSAID), carbamazepine (anticonvulsant and mood stabilizer), sulfamethoxazole (sulfonamide), furosemide (diuretic), and fenofibrate (lipid-lowering drug)—under MAE has been proposed in [133]. The method achieved an average recovery of $91.1\%$, while corresponding limits of detection (LOD) for the abovementioned pharmaceuticals were $9.23$, $0.88$, $3.61$, $35.68$, and $0.69$ ng/g, respectively. In [104], selected pharmaceuticals were detected in sediments along the Umgeni River. Therein, samples were analyzed using High-Performance Liquid Chromatography (HPLC) with Diode Array Detection (DAD) coupled with Electrospray Ionization Mass Spectrometry (ESI-MS), dubbed HPLC-DAD-ESI-MS, prior to ultrasonication and centrifugation, clean up and pre-concentration by solid-phase extraction (SPE). High analyte recoveries ranging within 71.20–118% were achieved for most of the considered analytes.

Ultrasonic-assisted extraction (UAE) coupled with high-performance liquid chromatography has also been widely employed for the determination of pharmaceuticals in sediment matrices. A representative study by Lei et al. [105] applied this approach to antibiotics in lyophilized lake sediments using a 30 mL mixture of acetonitrile and 0.1 M EDTA–McIlvaine buffer (1:1, v/v), followed by HPLC–ESI–MS/MS determination. Reported recoveries ranged from $72.5\%$ to $113.5\%$, with limits of detection between $0.11$ and $1.15$ $\mathsf{\mu }$g/kg. These findings demonstrate that ultrasonic-assisted extraction, when combined with optimized solvent systems and mass-spectrometric detection, can achieve efficient analyte desorption and satisfactory sensitivity for multiclass PPCPs in sediment samples.

Another modern extraction approach in analytical chemistry which derives from a modification of Solid-Phase Extraction (SPE) is Molecularly Imprinted Solid-Phase Extraction (MISPE). MISPE has been utilized in the extraction of pharmaceutical compounds from various sample matrices, but its specific application to pharmaceutical sediments is relatively limited. It utilizes magnetic particles functionalized with ion exchange or chelating agents for selective extraction. The incorporation of magnetic particles improves the efficiency and convenience of the extraction process compared to traditional SPE. Thus, MISPE offers several advantages in sample preparation and analysis related but not limited to selectivity, improved sensitivity, wide applicability/versatility, reusability, lower matrix interference and environmental friendliness.

A sensitive and effective method [106] was developed and validated for selective adsorption and quantitation of norfloxacin (NFX) from marine sediments, using novel molecularly imprinted silica polymers as sorbents, followed by high-performance liquid chromatographic analysis with diode array detection (HPLC-DAD). Several parameters affecting the extraction efficiency of MISPE were optimized. As a matter of fact, the obtained recoveries lied within 75.5–91.7% whereas LOD was 5 $\mathsf{\mu }$g/kg.

Parabens are synthetic preservatives used in cosmetics, personal care products, and some pharmaceutical formulations. Their purpose is to prevent microbial growth and extend product shelf life. Although parabens can be present in certain pharmaceuticals, they are not classified as pharmaceuticals themselves. Nevertheless, parabens and bisphenols have received attention due to their potential health effects, including their ability to mimic or interfere with hormone function in the body. They are considered endocrine-disrupting compounds (EDCs) and have been a subject of regulatory scrutiny and research regarding their potential impacts on human health and the environment.

HPLC–MS/MS analysis has also been applied for detecting parabens and bisphenols [134] as well as other PPCPs [121] in freeze-dried marine sediments. In the former approach, an off-line SLE method was employed, with the first extraction using 7 mL of methanol, and the second requiring 7 mL of ethyl acetate, followed by centrifugation. In the latter work, four solvents—MeOH (methanol), EtOH (ethanol), EtAc (ethyl acetate), and ACN (acetonitrile)—were tested in combination with 0.25 g of C18 material as a solid support, in vortex-assisted matrix solid-phase dispersion extraction (VA-MSPD). Recoveries reported in [134] ranged from $53\%$ to $112\%$ for matrix spikes, while Soares et al. [121] obtained average recoveries between $60\%$ and $140\%$ for $64\%$ of the compounds.

Regarding [121], the recovery efficiency of each solvent was evaluated. Although ACN and EtAc had similar polarities, they showed poor recovery for some compounds. Average recoveries were below $70\%$ for approximately $60\%$ of the analytes in the case of ACN as extraction solvent. EtAc on the other hand, which is commonly used in sample preparation procedures due to its non-mutagenic and non-bioaccumulative nature, resulted in co-extraction of many matrix components. As another indicative factor of the effect of physicochemical properties on the extraction performance, more polar compounds, especially those with low $log{K}_{\mathrm{ow}}$ values (<3), were not successfully extracted. When MeOH was used as the extraction solvent, it yielded the best extraction efficiency. MeOH’s effectiveness as an extraction solvent can be attributed to its ability to interact with polar compounds due to its polar protic nature. MeOH has a high dielectric constant ($ϵ$), indicating a higher probability of interaction with polar analytes through hydrogen bonding. Dielectric constant is an important parameter to assess solvent polarity.

In the case of extracting organic compounds from various sample matrices, including environmental, food, and biological samples, stir bar sorptive extraction (SBSE) is a another effective, commonly used technique. It is a solventless technique that uses a coated stir bar to extract analytes from a sample matrix. SBSE is valued for its simplicity, user-friendliness, and ability to concentrate analytes from complex samples. However, its application for the extraction of pharmaceuticals in sediments is not as common because of the potential interference from the complex matrix. Pharmaceutical sediments often contain excipients, fillers, and other components that can affect the extraction efficiency and the sorption properties of the stir bar. Therefore, other extraction techniques that are better suited for the specific characteristics of pharmaceutical sediments are often preferred.

To improve the effectiveness of the SBSE extraction process, Hu et al. [110] suggested that various parameters need to be studied and optimized. These include the extraction temperature and duration, stirring speed, pH level, ionic strength, desorption solvent and time. Their goal was to determine the optimal conditions that allow for high extraction efficiency. The authors proposed a new approach dubbed PANi/MWCNTs-OH-SBSE, combined with HPLC–UV for the simultaneous determination of polar, semi-polar and apolar compounds (phenols, Non-steroidal anti-inflammatory drugs (NSAIDs) and PCBs) in environmental samples. NSAIDs are non-volatile compounds, meaning they do not readily evaporate at room temperature. Actually, the extraction temperature had a minimal effect on the extraction efficiency of NSAIDs. This indicates that the extraction of NSAIDs was less influenced by temperature variations. The developed method exhibited several benefits, such as low limits of detection, high extraction efficiency, plus a wide linear range.

In [135] three classes of pharmaceuticals, non-steroidal anti-inflammatory drugs (NSAID) (Ibuprofen, paracetamol, diclofenac) that are used for treating pain, fever and inflammation, antibiotics (trimethoprim), as well as, antidepressants (citalopram) were determined in freeze-dried river sediments. An improved SPE method was developed employing MeOH and Ultraperformance™Liquid Chromatography (UPLC) with Electrospray Ionization (ESI) and a hybrid quadrupole-linear ion trap (QqLIT) mass spectrometer for tandem mass spectrometry (MS/MS) analysis. In this case, satisfactory recoveries ranging from $20\%$ to $86\%$ were achieved.

It is interesting to notice that only a few analytical methodologies were based on GC to determine pharmaceuticals [113,114,120,136]. This limited use of gas chromatography for pharmaceuticals can be attributed to a number of factors such as the volatility of analytes (pharmaceutical sediments often contain non-volatile or high-molecular-weight compounds that may not readily volatilize under the conditions required for GC), matrix interference (complex matrices containing excipients, fillers, and other non-volatile components can interfere with GC, or co-elute with the analytes of interest, leading to poor chromatographic resolution and inaccurate quantification), thermal stability (GC utilizes high temperatures for analyte vaporization and separation, though certain pharmaceutical sediments may contain heat-sensitive compounds that can degrade or alter, making GC unsuitable for their analysis), and analyte polarity (GC is ineffective for some pharmaceutical sediments which may contain polar or ionizable compounds and thus techniques like LC or ion chromatography (IC) may be more suitable).

In [113], ultrasonicated and centrifuged procedures have been employed whereas acetone-ethyl acetate was prepared as the extraction solvent for the solid sample. EDCs (BPA and NP) were analyzed using a GC-MS system to obtain respective recoveries $95.1\%$ and $93.6\%$. Reported limits of detection (LOD) for BPA and NP were $0.01$ and $0.1$ ng/L, respectively. 2 typical polycyclic musks (PCMs), namely, 1,3,4,6,7,8-hexahydro-4,6,6,7,8,8-hexamethylcyclopenta-(g)-2-benzopyran (HHCB) and 7-acetyl-1,1,3,4,4,6-hexamethyl-1,2,3,4-tetrahydronaphthalene (AHTN), have been detected by ultrasonication followed by (GC) with a Mass Selective Detector (MSD) in [120] with reported recoveries $82\%$ and $81\%$ for HHCB and AHTN, respectively.

8 PPCPs (ibuprofen, 2-benzyl-4-chlorophenol, naproxen, triclosan, ketoprofen, diclofenac, bisphenol A and estrone) were extracted using UAE–SPME–GC–MS in [136]. This method is rather environmentally friendly since UAE was performed using water at pH 3 with $1\%$ MeOH. In addition, the study reported good recoveries with minimal matrix effects, so the use of internal standards was not necessary.

SWE-DLLME, also known as Solid-Phase Microextraction Dispersive Liquid–Liquid Microextraction, is a sample preparation approach that combines the principles of SPME and Dispersive Liquid–Liquid Microextraction (DLLME). This technique offers improved sensitivity, selectivity and efficiency in the analysis of target analytes from complex sample matrices. Compared with widely-used techniques, such as PLE, SWE-DLLME typically uses smaller volumes of solvents, making it a more eco-friendly technique. However, it offers moderate extraction efficiency since it relies on the affinity of analytes to the SPME fiber coating and the partitioning between the fiber coating and the organic extraction solvent. On the contrary, PLE provides higher extraction yields due to the combination of high pressure and elevated temperature.

SWE-DLLME has been employed to determine 13 EDCs followed by GC-MS achieving satisfactory recoveries from $42.3\%$ (dienestrol) to $131.3\%$ (4,5a-dihydrotestosterone), except for diethyl stilbestrol ($15.0\%$) and nonylphenols ($29.8\%$). Reported limits of detection (LOD) ranged from $0.006$ ng/g to $0.639$ ng/g [114].

Table 1. PPCPs and their derivatives in sediments.

Analytes	Samples/Pre-Treatment	Extraction Method	Conditions (Extraction Process)	Determination	Recovery/LOD	Year	Ref.
13 antibiotics: AZI, ERY, CLA	River sediments/Air-dried	PLE	$0.2\%$ ${\mathrm{NH}}_3$ in MeOH	LC-MS/MS	80–100%, except for ERY-EE ($54\%$)	2021	[31]
19 antibiotics (ABs)	River sediments/freeze-dried	PLE, Clean-up by SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	LC-MS/MS	45–125%	2022	[94]
20 Illicit drugs	River sediments/freeze-dried	PLE, clean up by SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (9:1, v/v), 1250 psi at 50 °C for 5 min	LC-MS/MS	90–135%, LOD $\le 1.1$ ng/g (d.w.), except cannabinoid $0.84-2.3$ ng/g (d.w.)	2021	[32]
Anti-bacterial–triclocarban	River sediments/freeze-dried	PLE	$5\mathrm{mL}$ $\mathrm{MeOH}:$ ${\mathrm{H}}_2\mathrm{O}$ (60:40, v/v)	LC–HRMS	$89\%$, LOD in $0.01$–$0.1$ ng/g (d.w.)	2015	[115]
16 antibiotics, anticonvulsants, NSAIDs, analgesics, stimulants, antibacterial, antifungal agents, insect repellents, hormones	Surface marine sediment/freeze-dried	PLE, clean-up by SPE	$5\mathrm{mL}$ $\mathrm{MeOH}:$ $\mathrm{ultrapure}\mathrm{water}$ (1:1, v/v)	LC–HRMS	60–$100\%$, LOD in $0.1$–$0.3$ ng/g, LOQ in $0.3$–$0.9$ ng/g	2024	[107]
13 pharmaceuticals, 5 personal care products	Lake sediments/freeze-dried	PLE, clean-up by SPE	80 °C; $2\times$ MeOH and $\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:2, v/v)	LC–QqQ–MS	$12.6$–$108.1\%$, LOD in $0.01$–$3.75$ ng/g, LOQ in $0.04$–$12.51$ ng/g	2024	[122]
36 out of 56 PPCPs (antibiotic, analgesic, antiarrhythmic, antiepileptic, antilipidemic, antitumor, blood vessel dilator, and others)	River sediments	PLE, clean-up by SPE using Oasis HLB cartridges	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v with $0.5\%$ (v/v) ${\mathrm{NH}}_3\left(\mathrm{aq}\right)$, pH = 11)	LC–MS/MS	$85.7$–$124.6\%$	2020	[96]
Pharmaceutically active compounds (PhACs): carbamazepine (anticonvulsant and mood stabilizer), ciprofloxacin (antibiotic), sulfamethoxazole (antibiotic)	Lagoon sediments/lyophilized	PLE, clean-up by SPE using Oasis HLB cartridges	MeOH, 60 °C, 3 cycles of 5 min each, pre-heating at 70 °C for 5 min	LC–MS/MS	Carbamazepine: $\mathrm{LOD}=0.25$ ng/g, $\mathrm{LOQ}=1$ ng/g; Ciprofloxacin: $\mathrm{LOD}=0.5$ ng/g, $\mathrm{LOQ}=2$ ng/g	2024	[123]
Chiral drugs: $\beta$ -blockers, antidepressants, $\beta$-agonist, antihistamine, stimulants	River sediments/freeze-dried and sieved	PLE, Clean-up by SPE (Oasis HLB)	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v), 100 °C, 1500 psi, 2 cycles	LC-MS/Chiral-V enantioselective column	from $22\pm 3\%$ (R(-)-chlorpheniramine) to $93\pm 5\%$ (acebutolol-E1)	2020	[116]
31 pharmaceuticals (NSAIDs, sulfonamides, $\beta$-blockers, psychotropic drugs, hormones)	Marine sediments/freeze-dried	MAE, clean-up by SPE	400 W, 60 °C, 10 min; (i) 10 mL 10% $\mathrm{Mg}{\left({\mathrm{NO}}_3\right)}_2$ + sat. ${\mathrm{NH}}_4\mathrm{Cl}$ (9:1, v/v) + 0.5 mL MeOH; (ii) 10 mL MeOH:DCM (4:1, v/v), acidic pH (non-polar analytes)	LC–MS/MS	<20% to ∼$80\%$, LOD in $0.067$–$2.67\mathrm{ng}/\mathrm{mL}$, LOQ in $0.2$–$8.0\mathrm{ng}/\mathrm{mL}$	2025	[124]
Veterinary medicine: anthelmintic drugs (ADs), Flubendazole (FLU) and fenbendazole (FEN)	River sediments/air-dried, grounded and sieved	MAE	30 mL hexane:acetone (1:1, v/v) at 115 °C for 10 min, 400 W	LC-MS/MS	98.3–103.4%	2015	[117]
21 pharmaceuticals	River sediments/freeze-dried	UAE and centrifugation, clean-up by SPE	2.5 mL 1 M NaCl, 2.5 mL 1 M oxalic acid and 5 mL ethanol	LC-MS/MS	From $78\pm 3\%$ to $117\pm 3\%$, LOD in 1.29–7.20 ng/g	2023	[125]
6 pharmaceuticals	River sediments/freeze-dried	UAE, clean-up by SPE	$10\mathrm{mL}$ with $2\%$ ${\mathrm{NH}}_4\mathrm{OH}$ in MeOH	LC–MS/MS	$40.1$–$120.6\%$, LOD in $0.02$–$2.5$ ng/g	2018	[126]
11 abused drugs (amphetamine, METH, heroin, ketamine, ephedrine, cocaine, codeine, methadone, morphine, benzoylecgonine, methcathinone)	River sediments/freeze-dried and sieved	UAE and centrifugation	10 mL McIlvain buffer:MeOH (1:1, v/v)	LC-MS/MS	68–101%, LOD in 0.20–1.50 ng/g	2019	[108]
22 out of 28 antibiotics	River sediments/freeze-dried, ground, sieved	UAE & centrifugation	10 mL ACN:citric acid buffer, pH = 3, (1:1, v/v)	LC-MS/MS	53–149%	2020	[95]
12 antimicrobials	Dam sediments/lyophilization and sieving	UAE, clean-up by on-line SPE	ACN:Citrate Buffer, pH = 3	LC-MS/MS	89–119%, LOD in 0.40–5.1 $\mathsf{\mu }$g/kg	2016	[66]
32 antibiotics	River sediments/freeze-dried	UAE, clean-up by SPE	McIlvaine buffer:ACN (1:1, v/v) and $\mathrm{Mg}{\left({\mathrm{NO}}_3\right)}_2$-${\mathrm{NH}}_3$·${\mathrm{H}}_2$O (96:4, v/v)	LC-MS/MS	40–127%, (except enrofloxacin and marbofloxacin) LOD in 0.01–0.45 ng/g (d.w.)	2015	[97]
Antibiotic and antiretroviral drug cocktails (ARVDs)	River sediments/dried	UAE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (4:1, v/v)	LC-ESI-MS/MS	84.3–111.3%	2020	[98]
41 drugs of abuse and metabolites	River sediments/lyophilized	UAE, Clean-up by SPE, MeOH & MeOH–DCM elution	$10\mathrm{mL}$ ${\mathrm{H}}_2\mathrm{O}$:MeOH (1:1, v/v) pH 4.5 for 10 min	LC-MS/MS	≥$50\%$, $\mathrm{LOD}<1.32$ ng/g (d.w.)	2015	[23]
25 pharmaceutical compounds (14 antibiotics, 4 anti-epileptic and antidepressant drugs, 3 analgesic and anti-inflammatory agents, 3 beta-blockers, and 1 lipid regulator)	Lake sediments/freeze-dried	UAE, Clean-up by SPE	ACN & pH 3 citric acid buffer, 15 min	LC–MS/MS	70–111%, LOD in $0.3$–$3.2$ ng/kg, LOQ in $1.2$–$11.2$ ng/kg	2024	[127]
NSAIDs & anti-epileptic drugs	River sediment/air-dried	UAE, clean-up by SPE	20 mL ACN:MeOH (1:1, v/v)	LC-PDA	74–112%, LOD in 0.010–0.027 $\mathsf{\mu }$g/kg	2020	[109]
$2\mathrm{stimulants}$, $2\mathrm{antibiotics}$, $2\mathrm{antiepileptics}$, $4\mathrm{NSAIDs}$	River sediments	UAE	2 cycles acetone & 2 cycles MeOH, for 25 min	LC–PDA	69–$109\%$, LOD in $0.848$–$2.756$ $\mathsf{\mu }$g/kg, LOQ in $2.828$–$9.192$ $\mathsf{\mu }$g/kg	2025	[128]
85 PPCPs	Estuary sediments	QuEChERS, clean-up by PSA, Secondary Clean-up dSPE	20 mL ACN:$\mathrm{Milli}$-${\mathrm{QH}}_2\mathrm{O}$ (MQW, purified ${\mathrm{H}}_2\mathrm{O}$) (1:1, v/v)	LC-MS/MS	from $67\pm 9\%$ (sulfamethoxazole-13C6), to $108\pm 11\%$ (alprazolam-d5)	2022	[129]
Blood lipid regulators, analgesics, anti-inflammatory drugs and $\beta$-blockers	Lagoon sediments/freeze-dried	QuEChERS, clean up by dSPE	10 mL buffered ACN	LC-MS/MS	27–120%, LOD in 0.001–$0.75\mathrm{ng}/\mathrm{mL}$	2019	[91]
25 multiclass pharmaceuticals	River sediments/lyophilized	QuEChERS, clean-up by dSPE	10 mL ACN	LC-LIT/Orbitrap MS	64–101%	2019	[119]
EDCs	Marine sediment/air-dried, ground and sieved	Soxhlet, clean-up by SPE	$200\mathrm{mL}$ $\mathrm{MeOH}:$ acetone (1:1, v/v) for 8 h	LC-MS/MS	50.39–129.10%	2020	[16]
4 $\beta -lactam$ (BLs) and 4 polyether ionophore antibiotics (PEs)	River sediments	SPE	5 mL MeOH	LC-MS/MS	>$75\%$, except for AMOX	2015	[37]
40 antibiotics (cephalosporin, fluoroquinolone, lincosamide, macrolide, nitroimidazole, quinolone, sulfonamide and tetracycline groups)	River sediment/freeze-dried and homogenized	dSPE by using 100 mg mix of C18 and PSA (1:2, w/w) and 50 mg ${\mathrm{MgSO}}_4$	5 mL MeOH:ACN (1:3, v/v)	LC-MS/MS	24–162%, 48–151%, 51–159%, and 50–149% for 10, 20, 50 and 100 $\mathsf{\mu }$g/kg spiking levels, respectively	2020	[99]
19 anthelmintic drugs (ADs)	River sediments/air-dried	PLE, Clean-up by SPE	$3\times$ MeOH:${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v) at 70 °C, 100 bar, 5 min	UHPLC-ESI-MS/MS	31–90%	2020	[111]
34 pharmaceuticals	River sediments/freeze-dried, ground and sieved	UAE and centrifugation, clean-up by SPE with Oasis HLB cartridges	10 mL citrate buffer (pH 3): ACN (1:1, v/v)	UHPLC-MS/MS	43–118%, LOD in 0.01–0.6 ng/g (d.w.)	2019	[4]
7 Pharmaceuticals, 1 Personal Care Product and 3 Hormones	River sediments/dried	UAE, clean-up by SPE	5 mL MeOH→ 5 mL MeOH:${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)→ 2 mL acetone	UHPLC-MS/MS	54.0–94.4%	2015	[130]
4 pharmaceuticals: Anti-inflammatory (Ketorolac, Naproxen), Antibiotics (Ofloxacin, Ciprofloxacin), Anti-cancer (Ifosfamide, Cyclophosphamide), $\beta$-Blockers (Atenolol, Propranolol)	Less complex matrices of sediments	UAE	$4\mathrm{mL}$ EtOAc:MeOH (1:1, v/v)	UHPLC-MS/MS	87–113%, $\mathrm{LOD}=0.66$ ng/g	2015	[100]
111 organic micropollutants (OMPs), anti-epileptic, antineoplastic agents, antihistamine	Lake sediments/air-dried	UAE	(i) $4\mathrm{mL}$ $\mathrm{ACN}:$ ${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v), $0.1\%$ formic acid (FA), (ii) $4\mathrm{mL}$ ACN, 2-propanol, and ${\mathrm{H}}_2\mathrm{O}$ (3/3/4, v/v/v, $0.1\%$ FA)	UHPLC-MS/MS	81–104%	2020	[92]
25 PhACs in 6 categories: 9 antibiotics, 9 hormones, 3 NSAIDs, 2 antipsychotic drugs, 1 hypoglycemic drug, 1 antiviral drug	Marine sediments/freezed	UAE, clean-up by SPE	20 mL Citric acid buffer (pH 3):ACN (1:1, v/v)	UHPLC-ESI-MS/MS	61–117%	2020	[101]
22 EDCs	River and lake sediments/lyophilized and sieved	UAE, clean-up by SPE with GCB cartridge	10 mL MeOH:acetone (1:1, v/v)	UHPLC–MS	75–110%	2016	[112]
Pharmaceuticals	Marine surface sediments/freeze-dried	UAE, clean-up by SPE using Oasis HLB cartridges	$\mathrm{MeOH}:\mathrm{ACN}$ (2:1, v/v) for 15 min at 25 °C	UPLC–QTOF–MS	75–$105\%$, LOD in $0.3$–$3.9$ ng/g	2025	[131]
30 PPCPs	River sediments/freeze-dried	UAE, clean-up by SPE	$10\mathrm{mL}$ $\mathrm{ACN}:$ $\mathrm{deionized}\mathrm{water}$ (1:1)	UPLC–Orbitrap/MS	74–$109\%$, LOD: $0.02$–$1.21$ ng/g, LOQ: $0.5$–$14.5$ ng/g	2024	[132]
21 Psychiatric drugs (Carbamazepine, citalopram, fluoxetine, sertraline, trazodone, and venlafaxine) and 6 antibiotics (azithromycin, ciprofloxacin, clarithromycin, moxifloxacin, ofloxacin, and trimethoprim)	River sediments	QuEChERS, clean-up by SPE	0.5 mL of ACN: ultra-pure water (UPW) (3:7, v/v)	UHPLC-MS/MS	Psychiatric drugs: from $47.2\%$ (sertraline) to $104\%$ (diazepam). Sulfonamide and fluoroquinolone antibiotics: <$30\%$ and <$20\%$, respectively.	2020	[102]
32 PPCPs	Lagoon and estuary sediments/lyophilized	2 × SPE: (i) HLB cartridges activated with SDS, (ii) mixed HLB-cation exchange cartridges	12 mL MeOH: $\mathrm{Milli}$-${\mathrm{QH}}_2\mathrm{O}$ (1:1, v/v)	UHPLC-MS/MS	$60\%$ of the compounds had recoveries >$80\%$, $\mathrm{LOD}=10$ ng/g	2020	[86]
Cardiovascular drugs (atorvastatin, fenofibrate, bezafibrate), anticonvulsant and mood stabilizers (Carbamazepine), benzodiazepine medication (diazepam), opioid medications (codeine, morphine)	River sediments/freeze-dried	SLE, cleaned up by stir-disc SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	UHPLC-MS/MS	50–111%, LOD in 0.02–9.9 ng/g	2020	[118]
30 pharmaceuticals	Marine sediments	SLE	$4.5\mathrm{mL}$ ACN + $0.5\mathrm{mL}$ ${\mathrm{H}}_2\mathrm{O}$ (9:1, v/v)	UHPLC–TOF–MS	$74.3$–$115.4\%$, LOD: $0.02$–$0.23$ ng/g, LOQ: $0.06$–$0.69$ ng/g	2025	[65]
46 PPs: Antibiotic, anti-inflammatory and cardiovascular	Marine sediments	PLE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	HPLC-MS	>$75\%$	2020	[103]
NSAIDs, anticonvulsants, sulfonamides, diuretics, lipid-lowering drugs	Pond sediments/freeze-dried and grounded	MAE	20 mL of 35/35/30 of MeOH/acetone/miliQ	HPLC-MS	$91.1\%$, LOD in 0.69–35.68 ng/g	2019	[133]
Pharmaceuticals (antibiotics, antipyretics, stimulant, antiepileptic and antipsychotic drugs)	River sediments	UAE and centrifugation, Clean-up by SPE	$2\times 50$ mL MeOH, 50 mL acetone: Acetic acid (20:1, v/v) and 50 mL EtOAc	HPLC-DAD-ESI-MS	71.20–118%, LOD in 0.0006–0.7986 ng/g	2015	[104]
12 Antibiotics	Lake sediments/lyophilized	UAE & Centrifugation	30 mL (ACN: $0.1M$ EDTA-Mcilvaine buffer, (1:1, v/v)	HPLC–ESI MS/MS	72.5–113.5%, LOD in 0.11–1.15 $\mathsf{\mu }$g/kg	2015	[105]
Fluoroquinolone antibiotics (NFX)	Marine sediments/dried	Molecularly imprinted solid-phase extraction (MISPE)	10 mL DCM, sonicated 20 min, centrifuged 5 min at 4000 rpm	HPLC-DAD	75.5–91.7%, $\mathrm{LOD}=5$ $\mathsf{\mu }$g/kg	2020	[106]
Preservatives-parabens (PCPs)	Marine sediments/freeze-dried	off-line SLE/SPE Clean-up with Oasis MCX	(i) 7 mL MeOH for 60 min, (ii) 7 mL of EtOAc	HPLC-MS/MS	57–105% (blanks), 53–112% (spiked)	2019	[134]
Pharmaceuticals and personal care products (PPCPs)	Marine sediments/freeze-dried	VA-MSPD	MeOH	HPLC-MS/MS	60–140%, LOD in 0.13–5.70 ng/g	2021	[121]
Polar, semi-polar, apolar compounds (NSAIDs & others)	River sediments/dried	stir bar sorptive extraction (SBSE)	Aqueous SBSE (no solvent); desorption with MeOH	HPLC–UV	81.8–121.3%, LOD in 0.09–0.81 g/L	2015	[110]
5 PPs: NSAIDs, antibiotics and antidepressants	River sediments/freeze-dried	SPE	5 mL $100\%$ MeOH followed by 5 mL deionized water at a rate of 1 mL/min	Ultraperformance TM-ESI-(QqLIT) MS/MS	Ibuprofen $73\%$, Paracetamol $86\%$, Diclofenac $20\%$, Trimethoprim $63\%$, Citalopram $43\%$	2019	[135]
EDCs (such as bisphenol A (BPA) and nonylphenol (NP)	Lake sediments/lyophilized and sieved	UAE	5 mL Acetone:EtOAc	GC-MS	BPA:$95.1\%$, NP: $93.6\%$, LOD = $0.01$ (BPA) and $0.1$ (NP) ng/L	2019	[113]
8 PPCPs	River sediments/lyophilized	UAE–SPME	7 mL deionized ${\mathrm{H}}_2\mathrm{O}$ (pH 3) & $1\%$ MeOH	GC–MS	56–108%, $\mathrm{LOD}<0.25$ ng/g	2017	[136]
2 PCMs: HHCB and AHTN	River sediments/freeze-dried	UAE & centrifugation	50 mL of MeOH	GC-MSD	HHCB $82\pm 11\%$, AHTN $81\pm 9\%$	2020	[120]
13 EDCs	River sediments/freeze-dried and sieved	SWE-DLLME	Chlorobenzene (CBz) for DLLME and $10\%$ acetone for SWE at 150 °C	GC–MS	$42.3\%$ (dienestrol), $131.3\%$ (4,5$\alpha$-dihydrotestosterone), except for diethyl stilbestrol ($15\%$) and nonylphenols ($29.8\%$), LOD in 0.006–0.639 ng/g	2015	[114]

Table 1. PPCPs and their derivatives in sediments.

Analytes	Samples/Pre-Treatment	Extraction Method	Conditions (Extraction Process)	Determination	Recovery/LOD	Year	Ref.
13 antibiotics: AZI, ERY, CLA	River sediments/Air-dried	PLE	$0.2\%$ ${\mathrm{NH}}_3$ in MeOH	LC-MS/MS	80–100%, except for ERY-EE ($54\%$)	2021	[31]
19 antibiotics (ABs)	River sediments/freeze-dried	PLE, Clean-up by SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	LC-MS/MS	45–125%	2022	[94]
20 Illicit drugs	River sediments/freeze-dried	PLE, clean up by SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (9:1, v/v), 1250 psi at 50 °C for 5 min	LC-MS/MS	90–135%, LOD $\le 1.1$ ng/g (d.w.), except cannabinoid $0.84-2.3$ ng/g (d.w.)	2021	[32]
Anti-bacterial–triclocarban	River sediments/freeze-dried	PLE	$5\mathrm{mL}$ $\mathrm{MeOH}:$ ${\mathrm{H}}_2\mathrm{O}$ (60:40, v/v)	LC–HRMS	$89\%$, LOD in $0.01$–$0.1$ ng/g (d.w.)	2015	[115]
16 antibiotics, anticonvulsants, NSAIDs, analgesics, stimulants, antibacterial, antifungal agents, insect repellents, hormones	Surface marine sediment/freeze-dried	PLE, clean-up by SPE	$5\mathrm{mL}$ $\mathrm{MeOH}:$ $\mathrm{ultrapure}\mathrm{water}$ (1:1, v/v)	LC–HRMS	60–$100\%$, LOD in $0.1$–$0.3$ ng/g, LOQ in $0.3$–$0.9$ ng/g	2024	[107]
13 pharmaceuticals, 5 personal care products	Lake sediments/freeze-dried	PLE, clean-up by SPE	80 °C; $2\times$ MeOH and $\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:2, v/v)	LC–QqQ–MS	$12.6$–$108.1\%$, LOD in $0.01$–$3.75$ ng/g, LOQ in $0.04$–$12.51$ ng/g	2024	[122]
36 out of 56 PPCPs (antibiotic, analgesic, antiarrhythmic, antiepileptic, antilipidemic, antitumor, blood vessel dilator, and others)	River sediments	PLE, clean-up by SPE using Oasis HLB cartridges	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v with $0.5\%$ (v/v) ${\mathrm{NH}}_3\left(\mathrm{aq}\right)$, pH = 11)	LC–MS/MS	$85.7$–$124.6\%$	2020	[96]
Pharmaceutically active compounds (PhACs): carbamazepine (anticonvulsant and mood stabilizer), ciprofloxacin (antibiotic), sulfamethoxazole (antibiotic)	Lagoon sediments/lyophilized	PLE, clean-up by SPE using Oasis HLB cartridges	MeOH, 60 °C, 3 cycles of 5 min each, pre-heating at 70 °C for 5 min	LC–MS/MS	Carbamazepine: $\mathrm{LOD}=0.25$ ng/g, $\mathrm{LOQ}=1$ ng/g; Ciprofloxacin: $\mathrm{LOD}=0.5$ ng/g, $\mathrm{LOQ}=2$ ng/g	2024	[123]
Chiral drugs: $\beta$ -blockers, antidepressants, $\beta$-agonist, antihistamine, stimulants	River sediments/freeze-dried and sieved	PLE, Clean-up by SPE (Oasis HLB)	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v), 100 °C, 1500 psi, 2 cycles	LC-MS/Chiral-V enantioselective column	from $22\pm 3\%$ (R(-)-chlorpheniramine) to $93\pm 5\%$ (acebutolol-E1)	2020	[116]
31 pharmaceuticals (NSAIDs, sulfonamides, $\beta$-blockers, psychotropic drugs, hormones)	Marine sediments/freeze-dried	MAE, clean-up by SPE	400 W, 60 °C, 10 min; (i) 10 mL 10% $\mathrm{Mg}{\left({\mathrm{NO}}_3\right)}_2$ + sat. ${\mathrm{NH}}_4\mathrm{Cl}$ (9:1, v/v) + 0.5 mL MeOH; (ii) 10 mL MeOH:DCM (4:1, v/v), acidic pH (non-polar analytes)	LC–MS/MS	<20% to ∼$80\%$, LOD in $0.067$–$2.67\mathrm{ng}/\mathrm{mL}$, LOQ in $0.2$–$8.0\mathrm{ng}/\mathrm{mL}$	2025	[124]
Veterinary medicine: anthelmintic drugs (ADs), Flubendazole (FLU) and fenbendazole (FEN)	River sediments/air-dried, grounded and sieved	MAE	30 mL hexane:acetone (1:1, v/v) at 115 °C for 10 min, 400 W	LC-MS/MS	98.3–103.4%	2015	[117]
21 pharmaceuticals	River sediments/freeze-dried	UAE and centrifugation, clean-up by SPE	2.5 mL 1 M NaCl, 2.5 mL 1 M oxalic acid and 5 mL ethanol	LC-MS/MS	From $78\pm 3\%$ to $117\pm 3\%$, LOD in 1.29–7.20 ng/g	2023	[125]
6 pharmaceuticals	River sediments/freeze-dried	UAE, clean-up by SPE	$10\mathrm{mL}$ with $2\%$ ${\mathrm{NH}}_4\mathrm{OH}$ in MeOH	LC–MS/MS	$40.1$–$120.6\%$, LOD in $0.02$–$2.5$ ng/g	2018	[126]
11 abused drugs (amphetamine, METH, heroin, ketamine, ephedrine, cocaine, codeine, methadone, morphine, benzoylecgonine, methcathinone)	River sediments/freeze-dried and sieved	UAE and centrifugation	10 mL McIlvain buffer:MeOH (1:1, v/v)	LC-MS/MS	68–101%, LOD in 0.20–1.50 ng/g	2019	[108]
22 out of 28 antibiotics	River sediments/freeze-dried, ground, sieved	UAE & centrifugation	10 mL ACN:citric acid buffer, pH = 3, (1:1, v/v)	LC-MS/MS	53–149%	2020	[95]
12 antimicrobials	Dam sediments/lyophilization and sieving	UAE, clean-up by on-line SPE	ACN:Citrate Buffer, pH = 3	LC-MS/MS	89–119%, LOD in 0.40–5.1 $\mathsf{\mu }$g/kg	2016	[66]
32 antibiotics	River sediments/freeze-dried	UAE, clean-up by SPE	McIlvaine buffer:ACN (1:1, v/v) and $\mathrm{Mg}{\left({\mathrm{NO}}_3\right)}_2$-${\mathrm{NH}}_3$·${\mathrm{H}}_2$O (96:4, v/v)	LC-MS/MS	40–127%, (except enrofloxacin and marbofloxacin) LOD in 0.01–0.45 ng/g (d.w.)	2015	[97]
Antibiotic and antiretroviral drug cocktails (ARVDs)	River sediments/dried	UAE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (4:1, v/v)	LC-ESI-MS/MS	84.3–111.3%	2020	[98]
41 drugs of abuse and metabolites	River sediments/lyophilized	UAE, Clean-up by SPE, MeOH & MeOH–DCM elution	$10\mathrm{mL}$ ${\mathrm{H}}_2\mathrm{O}$:MeOH (1:1, v/v) pH 4.5 for 10 min	LC-MS/MS	≥$50\%$, $\mathrm{LOD}<1.32$ ng/g (d.w.)	2015	[23]
25 pharmaceutical compounds (14 antibiotics, 4 anti-epileptic and antidepressant drugs, 3 analgesic and anti-inflammatory agents, 3 beta-blockers, and 1 lipid regulator)	Lake sediments/freeze-dried	UAE, Clean-up by SPE	ACN & pH 3 citric acid buffer, 15 min	LC–MS/MS	70–111%, LOD in $0.3$–$3.2$ ng/kg, LOQ in $1.2$–$11.2$ ng/kg	2024	[127]
NSAIDs & anti-epileptic drugs	River sediment/air-dried	UAE, clean-up by SPE	20 mL ACN:MeOH (1:1, v/v)	LC-PDA	74–112%, LOD in 0.010–0.027 $\mathsf{\mu }$g/kg	2020	[109]
$2\mathrm{stimulants}$, $2\mathrm{antibiotics}$, $2\mathrm{antiepileptics}$, $4\mathrm{NSAIDs}$	River sediments	UAE	2 cycles acetone & 2 cycles MeOH, for 25 min	LC–PDA	69–$109\%$, LOD in $0.848$–$2.756$ $\mathsf{\mu }$g/kg, LOQ in $2.828$–$9.192$ $\mathsf{\mu }$g/kg	2025	[128]
85 PPCPs	Estuary sediments	QuEChERS, clean-up by PSA, Secondary Clean-up dSPE	20 mL ACN:$\mathrm{Milli}$-${\mathrm{QH}}_2\mathrm{O}$ (MQW, purified ${\mathrm{H}}_2\mathrm{O}$) (1:1, v/v)	LC-MS/MS	from $67\pm 9\%$ (sulfamethoxazole-13C6), to $108\pm 11\%$ (alprazolam-d5)	2022	[129]
Blood lipid regulators, analgesics, anti-inflammatory drugs and $\beta$-blockers	Lagoon sediments/freeze-dried	QuEChERS, clean up by dSPE	10 mL buffered ACN	LC-MS/MS	27–120%, LOD in 0.001–$0.75\mathrm{ng}/\mathrm{mL}$	2019	[91]
25 multiclass pharmaceuticals	River sediments/lyophilized	QuEChERS, clean-up by dSPE	10 mL ACN	LC-LIT/Orbitrap MS	64–101%	2019	[119]
EDCs	Marine sediment/air-dried, ground and sieved	Soxhlet, clean-up by SPE	$200\mathrm{mL}$ $\mathrm{MeOH}:$ acetone (1:1, v/v) for 8 h	LC-MS/MS	50.39–129.10%	2020	[16]
4 $\beta -lactam$ (BLs) and 4 polyether ionophore antibiotics (PEs)	River sediments	SPE	5 mL MeOH	LC-MS/MS	>$75\%$, except for AMOX	2015	[37]
40 antibiotics (cephalosporin, fluoroquinolone, lincosamide, macrolide, nitroimidazole, quinolone, sulfonamide and tetracycline groups)	River sediment/freeze-dried and homogenized	dSPE by using 100 mg mix of C18 and PSA (1:2, w/w) and 50 mg ${\mathrm{MgSO}}_4$	5 mL MeOH:ACN (1:3, v/v)	LC-MS/MS	24–162%, 48–151%, 51–159%, and 50–149% for 10, 20, 50 and 100 $\mathsf{\mu }$g/kg spiking levels, respectively	2020	[99]
19 anthelmintic drugs (ADs)	River sediments/air-dried	PLE, Clean-up by SPE	$3\times$ MeOH:${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v) at 70 °C, 100 bar, 5 min	UHPLC-ESI-MS/MS	31–90%	2020	[111]
34 pharmaceuticals	River sediments/freeze-dried, ground and sieved	UAE and centrifugation, clean-up by SPE with Oasis HLB cartridges	10 mL citrate buffer (pH 3): ACN (1:1, v/v)	UHPLC-MS/MS	43–118%, LOD in 0.01–0.6 ng/g (d.w.)	2019	[4]
7 Pharmaceuticals, 1 Personal Care Product and 3 Hormones	River sediments/dried	UAE, clean-up by SPE	5 mL MeOH→ 5 mL MeOH:${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)→ 2 mL acetone	UHPLC-MS/MS	54.0–94.4%	2015	[130]
4 pharmaceuticals: Anti-inflammatory (Ketorolac, Naproxen), Antibiotics (Ofloxacin, Ciprofloxacin), Anti-cancer (Ifosfamide, Cyclophosphamide), $\beta$-Blockers (Atenolol, Propranolol)	Less complex matrices of sediments	UAE	$4\mathrm{mL}$ EtOAc:MeOH (1:1, v/v)	UHPLC-MS/MS	87–113%, $\mathrm{LOD}=0.66$ ng/g	2015	[100]
111 organic micropollutants (OMPs), anti-epileptic, antineoplastic agents, antihistamine	Lake sediments/air-dried	UAE	(i) $4\mathrm{mL}$ $\mathrm{ACN}:$ ${\mathrm{H}}_2\mathrm{O}$ (1:1, v/v), $0.1\%$ formic acid (FA), (ii) $4\mathrm{mL}$ ACN, 2-propanol, and ${\mathrm{H}}_2\mathrm{O}$ (3/3/4, v/v/v, $0.1\%$ FA)	UHPLC-MS/MS	81–104%	2020	[92]
25 PhACs in 6 categories: 9 antibiotics, 9 hormones, 3 NSAIDs, 2 antipsychotic drugs, 1 hypoglycemic drug, 1 antiviral drug	Marine sediments/freezed	UAE, clean-up by SPE	20 mL Citric acid buffer (pH 3):ACN (1:1, v/v)	UHPLC-ESI-MS/MS	61–117%	2020	[101]
22 EDCs	River and lake sediments/lyophilized and sieved	UAE, clean-up by SPE with GCB cartridge	10 mL MeOH:acetone (1:1, v/v)	UHPLC–MS	75–110%	2016	[112]
Pharmaceuticals	Marine surface sediments/freeze-dried	UAE, clean-up by SPE using Oasis HLB cartridges	$\mathrm{MeOH}:\mathrm{ACN}$ (2:1, v/v) for 15 min at 25 °C	UPLC–QTOF–MS	75–$105\%$, LOD in $0.3$–$3.9$ ng/g	2025	[131]
30 PPCPs	River sediments/freeze-dried	UAE, clean-up by SPE	$10\mathrm{mL}$ $\mathrm{ACN}:$ $\mathrm{deionized}\mathrm{water}$ (1:1)	UPLC–Orbitrap/MS	74–$109\%$, LOD: $0.02$–$1.21$ ng/g, LOQ: $0.5$–$14.5$ ng/g	2024	[132]
21 Psychiatric drugs (Carbamazepine, citalopram, fluoxetine, sertraline, trazodone, and venlafaxine) and 6 antibiotics (azithromycin, ciprofloxacin, clarithromycin, moxifloxacin, ofloxacin, and trimethoprim)	River sediments	QuEChERS, clean-up by SPE	0.5 mL of ACN: ultra-pure water (UPW) (3:7, v/v)	UHPLC-MS/MS	Psychiatric drugs: from $47.2\%$ (sertraline) to $104\%$ (diazepam). Sulfonamide and fluoroquinolone antibiotics: <$30\%$ and <$20\%$, respectively.	2020	[102]
32 PPCPs	Lagoon and estuary sediments/lyophilized	2 × SPE: (i) HLB cartridges activated with SDS, (ii) mixed HLB-cation exchange cartridges	12 mL MeOH: $\mathrm{Milli}$-${\mathrm{QH}}_2\mathrm{O}$ (1:1, v/v)	UHPLC-MS/MS	$60\%$ of the compounds had recoveries >$80\%$, $\mathrm{LOD}=10$ ng/g	2020	[86]
Cardiovascular drugs (atorvastatin, fenofibrate, bezafibrate), anticonvulsant and mood stabilizers (Carbamazepine), benzodiazepine medication (diazepam), opioid medications (codeine, morphine)	River sediments/freeze-dried	SLE, cleaned up by stir-disc SPE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	UHPLC-MS/MS	50–111%, LOD in 0.02–9.9 ng/g	2020	[118]
30 pharmaceuticals	Marine sediments	SLE	$4.5\mathrm{mL}$ ACN + $0.5\mathrm{mL}$ ${\mathrm{H}}_2\mathrm{O}$ (9:1, v/v)	UHPLC–TOF–MS	$74.3$–$115.4\%$, LOD: $0.02$–$0.23$ ng/g, LOQ: $0.06$–$0.69$ ng/g	2025	[65]
46 PPs: Antibiotic, anti-inflammatory and cardiovascular	Marine sediments	PLE	$\mathrm{MeOH}:{\mathrm{H}}_2\mathrm{O}$ (1:1, v/v)	HPLC-MS	>$75\%$	2020	[103]
NSAIDs, anticonvulsants, sulfonamides, diuretics, lipid-lowering drugs	Pond sediments/freeze-dried and grounded	MAE	20 mL of 35/35/30 of MeOH/acetone/miliQ	HPLC-MS	$91.1\%$, LOD in 0.69–35.68 ng/g	2019	[133]
Pharmaceuticals (antibiotics, antipyretics, stimulant, antiepileptic and antipsychotic drugs)	River sediments	UAE and centrifugation, Clean-up by SPE	$2\times 50$ mL MeOH, 50 mL acetone: Acetic acid (20:1, v/v) and 50 mL EtOAc	HPLC-DAD-ESI-MS	71.20–118%, LOD in 0.0006–0.7986 ng/g	2015	[104]
12 Antibiotics	Lake sediments/lyophilized	UAE & Centrifugation	30 mL (ACN: $0.1M$ EDTA-Mcilvaine buffer, (1:1, v/v)	HPLC–ESI MS/MS	72.5–113.5%, LOD in 0.11–1.15 $\mathsf{\mu }$g/kg	2015	[105]
Fluoroquinolone antibiotics (NFX)	Marine sediments/dried	Molecularly imprinted solid-phase extraction (MISPE)	10 mL DCM, sonicated 20 min, centrifuged 5 min at 4000 rpm	HPLC-DAD	75.5–91.7%, $\mathrm{LOD}=5$ $\mathsf{\mu }$g/kg	2020	[106]
Preservatives-parabens (PCPs)	Marine sediments/freeze-dried	off-line SLE/SPE Clean-up with Oasis MCX	(i) 7 mL MeOH for 60 min, (ii) 7 mL of EtOAc	HPLC-MS/MS	57–105% (blanks), 53–112% (spiked)	2019	[134]
Pharmaceuticals and personal care products (PPCPs)	Marine sediments/freeze-dried	VA-MSPD	MeOH	HPLC-MS/MS	60–140%, LOD in 0.13–5.70 ng/g	2021	[121]
Polar, semi-polar, apolar compounds (NSAIDs & others)	River sediments/dried	stir bar sorptive extraction (SBSE)	Aqueous SBSE (no solvent); desorption with MeOH	HPLC–UV	81.8–121.3%, LOD in 0.09–0.81 g/L	2015	[110]
5 PPs: NSAIDs, antibiotics and antidepressants	River sediments/freeze-dried	SPE	5 mL $100\%$ MeOH followed by 5 mL deionized water at a rate of 1 mL/min	Ultraperformance TM-ESI-(QqLIT) MS/MS	Ibuprofen $73\%$, Paracetamol $86\%$, Diclofenac $20\%$, Trimethoprim $63\%$, Citalopram $43\%$	2019	[135]
EDCs (such as bisphenol A (BPA) and nonylphenol (NP)	Lake sediments/lyophilized and sieved	UAE	5 mL Acetone:EtOAc	GC-MS	BPA:$95.1\%$, NP: $93.6\%$, LOD = $0.01$ (BPA) and $0.1$ (NP) ng/L	2019	[113]
8 PPCPs	River sediments/lyophilized	UAE–SPME	7 mL deionized ${\mathrm{H}}_2\mathrm{O}$ (pH 3) & $1\%$ MeOH	GC–MS	56–108%, $\mathrm{LOD}<0.25$ ng/g	2017	[136]
2 PCMs: HHCB and AHTN	River sediments/freeze-dried	UAE & centrifugation	50 mL of MeOH	GC-MSD	HHCB $82\pm 11\%$, AHTN $81\pm 9\%$	2020	[120]
13 EDCs	River sediments/freeze-dried and sieved	SWE-DLLME	Chlorobenzene (CBz) for DLLME and $10\%$ acetone for SWE at 150 °C	GC–MS	$42.3\%$ (dienestrol), $131.3\%$ (4,5$\alpha$-dihydrotestosterone), except for diethyl stilbestrol ($15\%$) and nonylphenols ($29.8\%$), LOD in 0.006–0.639 ng/g	2015	[114]

4.2. Discussion on Our Findings

Although the present work primarily constitutes a systematic and methodological review, it also integrates a quantitative component derived from the authors’ own synthesis of data extracted from the 54 reviewed studies. This analysis, which preceded the initiation of ongoing laboratory work on PPCP determination at our research facilities, was designed to identify prevailing analytical trends and benchmark recovery and sensitivity performance across extraction families. The laboratory component of our project, still in progress, has been constrained by instrumental and sampling limitations; therefore, the current section focuses on the meta-analytical evaluation of published methods as an essential preparatory step toward future experimental optimization.

To provide a representative and statistically balanced overview of method performance, a concise meta-analysis was conducted using the subset of extraction techniques reported in at least two independent studies ($n\ge 2$). This filtering ensured fair comparison between methods and avoided bias from single-case reports, while maintaining coverage of the main extraction families summarized in Table 1. For each technique, the mean recovery and the median lower/upper bounds of the reported detection limits (LOD) were calculated. The results, illustrated in Figure 6, delineate the general performance envelopes of the principal extraction methods applied to PPCPs in sediments.

Overall, ultrasound- and pressure-assisted extractions (UAE, PLE, MAE) show the most favorable combination of high recoveries and low detection limits. Specifically, UAE (23 studies) exhibited a mean recovery of 88.9% and LOD in $0.065$–$1.265$ ng/g; PLE (11 studies) averaged 82.8% recovery with LOD in $0.10$–$0.50$ ng/g; MAE (3 studies) achieved 80.6% recovery and broader LOD in $0.378$–$19.175$ ng/g. Among other approaches, SLE (3 studies) reached 85.6% recovery with LOD in $0.020$–$5.065$ ng/g, QuEChERS (4 studies) 79.8% recovery and LOD in $0.001$–$0.750$ ng/g, whereas SPE-based methods (3 studies) produced recoveries around 70.7% and higher detection limits near 10 ng/g.

These trends indicate that UAE and PLE remain the most reproducible and analytically efficient techniques for PPCP determination in sediment matrices, offering recoveries close to 90% and sub-ng/g sensitivity under optimized conditions. Figure 6 quantitatively supports the evidence-based recommendations further discussed in Section 5.

5. Conclusions

This review provides a comprehensive overview of analytical methodologies for the detection of pharmaceuticals and personal care products in aquatic sediments. The discussion focused on literature from the past decade (2015–2025), emphasizing all stages of the analytical pipeline, from sample pre-processing and extraction to clean-up, instrumental analysis, and performance evaluation.

Building on the quantitative synthesis presented in Section 4.2 and Figure 6, the following conclusions summarize the key methodological trends and evidence-based recommendations for PPCP determination in sediment matrices.

We proposed a taxonomy of extraction-based workflows, with classification criteria including pre-treatment type, extraction technique, reported recoveries, and detection limits. Our findings show that UAE is by far the most frequently employed extraction approach for PPCPs in sediment matrices, representing 42.6% of the reviewed studies. It is often coupled with Solid-Phase Extraction (SPE) for post-extraction clean-up to mitigate matrix effects. Among the top employed methods, other widely used techniques include Pressurized Liquid Extraction (PLE, 20.4%), QuEChERS (7.4%), Microwave-Assisted Extraction (MAE, 5.6%), and Solid–Liquid Extraction (SLE, 5.6%), each selected according to analyte class, matrix characteristics, and available instrumentation.

Regarding instrumental determination, Liquid Chromatography (LC) and its high-performance variants (UPLC, HPLC) coupled with tandem mass spectrometry (MS/MS) dominate PPCP analysis due to the polar and thermally labile nature of these compounds. Gas Chromatography–Mass Spectrometry (GC–MS) is employed less frequently and typically requires derivatization to enhance volatility and sensitivity.

Across the reviewed studies, average recovery rates and limits of detection (LOD) served as key performance metrics. These outcomes reflect the interdependence of all steps in the analytical workflow, from solvent compatibility and extraction conditions to analyte-specific physicochemical behavior. Method optimization should therefore be tailored not only to the target compound classes but also to sediment characteristics, including grain size, organic-matter content, and matrix complexity.

Overall, our structured survey consolidates current knowledge and provides a clear methodological framework for selecting efficient, sensitive, and environmentally conscious extraction strategies for PPCP determination in sediments. By systematizing extraction workflows and linking them to performance outcomes, the proposed classification enhances reproducibility and supports the design of sustainable monitoring programs.

Future Research Agenda and Knowledge Gaps

To further advance PPCP analysis in complex sediment matrices, future research should address the following interrelated directions:

(i)

Harmonization and inter-laboratory comparability. The lack of harmonized reference procedures remains a major barrier in environmental-matrix analyses. Establishing validated, consensus protocols for extraction, clean-up, and quantification, along with structured inter-laboratory comparisons and personnel training, will improve data reliability and enable reproducible and comparable results across independent facilities.

(ii)

Transformation products (TPs) and real-matrix evaluation. A significant knowledge gap concerns transformation products: Approximately 40–45% of PPCPs lack TP information, and most available data derive from synthetic rather than real sediment matrices [137]. Future studies should prioritize matrix-realistic conditions, harmonized TP identification workflows, and toxicity evaluation of environmentally persistent by-products.

(iii)

Miniaturized and solvent-free extraction technologies. Advances in green analytical chemistry should focus on developing miniaturized, low-solvent, or solvent-free extraction systems, such as deep-eutectic-solvent- or $\mu$MSPD-based approaches. Emphasis should be placed on solvent reusability, energy efficiency, and quantitative greenness assessment to align PPCP analysis with sustainability principles.

Addressing these priorities will help bridge methodological fragmentation, improve comparability of sediment-monitoring data worldwide, and align PPCP analytics with the broader goals of sustainability and circular laboratory practice.