According to the most popular definition given by the United States Geological Survey, an emerging contaminant (EC) is “any synthetic or naturally occurring chemical that is not commonly monitored in the environment but has the potential to enter the environment and cause known or suspected adverse ecological and/or human health effects” [1
NORMAN Network Europe (network of reference laboratories, research centers, and related organizations for the monitoring and biomonitoring of emerging environmental substances) differentiates between emerging substances and emerging pollutants. Emerging substances are defined as “substances that have been detected in the environment, but which are currently not included in routine monitoring programs at EU level and whose fate, behavior, and (eco)toxicological effects are not well understood”. Emerging pollutants are defined as “pollutants that are currently not included in routine monitoring programs at the European level and which may be candidates for future regulation, depending on research on their (eco)toxicity, potential health effects and public perception and on monitoring data regarding their occurrence in the various environmental compartments” [3
The EC group contains substances which vary in regard to their chemical structure, toxicity, and environmental behavior and includes, among others, industrial additives and by-products, personal care products, and human and animal pharmaceuticals (PPCPs), surfactants, flame retardants, hormones, and sterols [3
]. This list is not complete and each year is extended with newly-detected artificial contaminants as well as naturally occurring trace compounds. In recent years it has expanded through the inclusion of nanomaterials and microplastic particles [4
]. More than 2100 scientific studies published between 2007 and 2016 have proven that ECs, as biologically active compounds, exhibit a potential risk to humans, plants, and/or animals [2
]. Many of them are able to alter the normal hormone function of wildlife and humans by mimicking or magnifying the effects of endogenous hormones, disrupting their synthesis and activity or the operation of hormone receptors. Strongest endocrine activity is demonstrated by natural and synthetic hormones which are introduced into the environment with insufficiently treated wastewater [5
]. Estrone (E1) and β-estradiol (E2) are two common forms of natural estrogen secreted by the human body which are frequently found in aqueous environments. This is especially true in respect to E2 due to its widespread use as a contraceptive and in hormone replacement therapy (HRT). Another non-steroidal synthetic estrogen, diethylstilbestrol (DES), has been formerly used medicinally to prevent stillbirths and as a growth stimulant in feed given to poultry, cattle, and sheep. Despite the fact that its effects have been proven harmful it is still used in the treatment of breast and prostate cancer and, in some countries, in HRT [7
]. Additionally, it has been shown that compounds making up some personal care products and industrial chemicals exhibit hormonal activity. The largest number of studies confirming their effect on the endocrine system is related to bisphenol A (BPA). It is suspected that estrogenic activity of BPA increases the risk of developing breast cancer in humans and may act as an antiandrogen causing feminizing side effects in men [9
]. Other ECs, including, for example, propylparaben (PP), butylparaben (BP), 3-(4-methylbenzylidene)camphor (3MBC), 4-t-octylphenol (4OP), 4-n-nonylphenol (4NP), and triclosan (TRC), have also been confirmed or suspected of having endocrine disrupting effects on the functioning of estrogen, androgen, prolactin, insulin, or thyroid hormones. It is supposed that continuous exposure to some ECs causes increased birth weight in children, adult fat gain, diabetes, and may potentially affect eating disorders [9
In recent years, micro-extraction in the liquid–liquid system (LLME) has become one of the most widely used techniques for the preparation of samples for EC determination [16
]. The most commonly used LLME modification is a dispersive liquid–liquid microextraction (DLLME), developed in 2006 by Rezaee et al. [18
]. This technique uses a ternary system consisting of an examined aqueous solution, an extraction solvent, and a dispersing solvent. The formation of the emulsion results in an unlimited contact area between two aqueous and organic phases producing prompt mass exchange. It is possible; however, to avoid the addition of a dispersing solvent to form the emulsion through the use of ultrasonic radiation. In 2008, Regueiro et al. [19
] used ultrasound for the first time to support microextraction in a liquid–liquid system, developing the ultrasound-assisted emulsification-microextraction technique (USAEME). In liquid–liquid microextraction techniques it is usual to use solvents which are heavier than water, although these substances are mostly chlorinated. This is due to the fact that after the extraction process the microdroplet of the organic solvent in which the analyte is dissolved is located at the bottom of the tube. Its collection (e.g., with a syringe) is relatively easy, especially if test tubes with a conical bottom are used. In case of solvents that are lighter than water, it is much more difficult to separate the microdroplet of the organic solvent from the aqueous phase. One solution which facilitates work with such solvents uses the process of solidifying the floating solvent drop. The technique using the process of solidification of the floating organic drop microextraction (SFODME) was introduced in 2007 by Khalili Zanjani et al. [20
]. In this technique, after the extraction process, the sample is placed in an ice bath to solidify the drop of the organic solvent in the upper part of the vessel in which the extraction is carried out. The drop is then transferred to a vial where it melts at room temperature. The solidification of the floating organic drop (SFOD) technique is combined with various liquid–liquid microextraction variants and used to determine both organic compounds and metal ions in various types of matrices [21
In this study, a simple and sensitive analytical procedure for simultaneous determination of hormones and other EC compounds most frequently detected in surface water bodies is optimized. USAEME-SFOD is used for the separation and preconcentration of analytes, whereas GC-MS in the selected ion monitoring (SIM) mode is applied for their quantification. The influence of extraction and derivatization parameters (i.e., the type of organic solvent and solvent volume, extraction time, derivatization reagent volume, and amount of buffering salt) on analyte recovery is investigated. The developed USAEME-SFOD/GC-MS procedure was used to assay target compounds in groundwater samples from northeastern Poland. To our knowledge, this is the first study utilizing USAEME with a non-chlorinated solvent for the determination of studied compounds. The presence of compounds from the EC group in groundwater has been largely unexplored, especially when compared to surface and marine waters, and this work may provide important knowledge within this area.
2.1. Reagents and Solvents
Materials, PP, BP, BPH, 3MBC, N,N-diethyltoluamide (DEET), OP, NP, TRC, BPA, diclofenac (DIC), E1, E2, DES, tricosane, and 1-undecanol, were obtained from Sigma-Aldrich (Darmstadt, Germany). Methanol and anhydrous disodium hydrogen phosphate (V) were provided by POCH (Gliwice, Poland). Acetic anhydride was purchased from Chempur (Piekary Śląskie, Poland). Stock solutions of each analyte (at 1 mg/mL of each) were prepared separately in methanol and stored at −18 °C for a period not exceeding one month. Working solutions were prepared by diluting the stock standard solution in methanol and storing them at −18 °C not longer than two weeks. Deionized water was obtained using a purification system (Milli-Q RG, Millipore, Burlington, MA, USA) and stored in glass bottles.
2.2. Groundwater Samples
Samples of groundwater were collected from six deep wells (the surface of water was at a depth of 15–46 m) and five shallow wells (the surface of the water was at a depth of 3–8 m) used for individual water supply. The wells are located in a region which is not directly affected by industrial sources of pollution. Samples of groundwater from contaminated sites (twelve) were collected from monitoring wells located in two municipal solid waste (MSW) landfill sites of non-hazardous and inert waste. All the sampling points were located in northeastern Poland. The geological structure of areas from which the samples were taken consists of clay–sand–gravel deposits. Samples were collected in glass bottles with Teflon-lined caps that were rinsed with the sample water on site and immediately carried to the laboratory where, upon arrival, they were filtered through a 0.45 µm pore size membrane filter and stored at −18 °C.
2.3. The Procedure of Ultrasound-Assisted Emulsification Microextraction with Solidification of Floating Organic Droplet (USAEME-SFOD) Coupled with In Situ Derivatization
For the simultaneous USAEME-SFOD and derivatization aliquots of 5 mL water samples were placed in 10 mL glass centrifuge tubes containing previously weighted 0.25 g of sodium hydrogen phosphate. The extraction solvent (1-undecanol, 20 µL) consisting of tricosane (5 µg/L) as an internal standard and the derivatization reagent (acetic anhydride, 250 µL) were added to the water sample and mixed. Immediately after, the tube was immersed in the ultrasonic Unitra Unima (Warsaw, Poland) water bath. Extractions were performed at 42 kHz ultrasound frequency and 230 W power for the duration of 8 min at room temperature. Emulsions were separated using centrifugation at 4000 rpm/min for 4 min in an MPW-250 Med. Instruments (Warsaw, Poland) laboratory centrifuge. After this process, the droplet of organic phase floated at the top of the test tube. The test tube was then cooled in an ice bath. After five minutes, 1-undecanol solidified and was transferred into a 150 µL micro vial with an integrated insert. It melted quickly at room temperature and GC-MS analysis of 1 µL of obtained solution was then performed as described in Section 2.4
. The course of the USAEME-SFOD is shown in Figure 1
2.4. GC-MS Conditions
Analysis was performed with an HP 6890 gas chromatograph coupled with a MSD5973 mass spectrometric detector and an HP 7673 autosampler (Agilent Technologies, Santa Clara, CA, USA). This device was equipped with an HP-5MS (5% phenylmethylsiloxane) column (size 30 m length × 0.25 mm; i.e., coated with 0.25 µm film thickness) and split/splitless injector. The injector was set to work in the splitless mode. Helium of 99.999% purity was used as a carrier gas at a flow rate of 1 mL/min. The injector temperature was set at 250 °C. The oven temperature program started from 160 °C and increased at increments of 2 °C/min to 170 °C (held for 2 min), 6.44 °C/min to 226 °C (held for 1 min), 10 °C/min to 233 °C (held for 2.5 min) and 10 °C/min to 300 °C (held for 4 min). The total run time was 26 min with the solvent delay time reaching 5.7 min. The MS detector worked in selected ion monitoring (SIM) mode. The electron impact source temperature was 230 °C with electron energy of 70 eV. The quadrupole temperature was 150 °C and the GC interface temperature was 280 °C. The chromatogram obtained during the GC-MS analysis of BPH, DEET, 3MBC, BPA, PP, BP, DES, DIC, OP, NP, E1, E2, and TRC together with the internal standard after USAEME-SFOD with in-situ acetylation is presented in Figure 2
. The MS spectra of target compounds are given in Figure S1 (Supplementary Material)
. The chromatographic parameters, molecular weights of target compounds, together with quantification and identification ions are shown in Table 1
. Based on the registered chromatograms, the relative areas of the chromatographic peaks were determined by dividing the values of the obtained peak areas of tested compounds by the peak area of the tricosane.