Next Article in Journal
Silver Nanoparticles Grown on Cross-Linked Poly (Methacrylic Acid) Microspheres: Synthesis, Characterization, and Antifungal Activity Evaluation
Next Article in Special Issue
Expediting Disulfiram Assays through a Systematic Analytical Quality by Design Approach
Previous Article in Journal
A High-Response Electrochemical As(III) Sensor Using Fe3O4–rGO Nanocomposite Materials
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Chemosensors
Volume 9
Issue 7
10.3390/chemosensors9070151
chemosensors-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Recent Advances in Solid-Phase Extraction (SPE) Based on Molecularly Imprinted Polymers (MIPs) for Analysis of Hormones

by
Anele Mpupa
1,2,
Shirley Kholofelo Selahle
1,2,
Boris Mizaikoff
1,3,* and
Philiswa Nosizo Nomngongo
1,2,4,*
1
Department of Chemical Sciences, Doornfontein Campus, University of Johannesburg, P.O. Box 17011, Johannesburg 2028, South Africa
2
Department of Science and Innovation (DSI)/National Research Foundation (NRF) South African Research Chair (SARChI), Nanotechnology for Water, University of Johannesburg, Doornfontein 2028, South Africa
3
Institute of Analytical and Bioanalytical Chemistry, Ulm University, Albert-Einstein-Allee 11, 89081 Ulm, Germany
4
DSI/Mintek Nanotechnology Innovation Centre, University of Johannesburg, Doornfontein 2028, South Africa
*
Authors to whom correspondence should be addressed.
Chemosensors 2021, 9(7), 151; https://doi.org/10.3390/chemosensors9070151
Submission received: 28 April 2021 / Revised: 17 June 2021 / Accepted: 18 June 2021 / Published: 22 June 2021
(This article belongs to the Collection Recent Trend in Chromatography for Pharmaceutical Analysis)
Browse Figures
Versions Notes

Abstract

:
Steroid hormones are active substances that are necessary in the normal functioning of all physiological activities in the body, such as sexual characteristics, metabolism, and mood control. They are also widely used as exogenous chemicals in medical and pharmaceutical applications as treatments and at times growth promoters in animal farming. The vast application of steroid hormones has resulted in them being found in different matrices, such as food, environmental, and biological samples. The presence of hormones in such matrices means that they can easily come into contact with humans and animals as exogenous compounds, resulting in abnormal concentrations that can lead to endocrine disruption. This makes their determination in different matrices a vital part of pollutant management and control. Although advances in analytical instruments are constant, it has been determined that these instruments still require some sample preparation steps to be able to determine the occurrence of pollutants in the complex matrices in which they occur. Advances are still being made in sample preparation to ensure easier, selective, and sensitive analysis of complex matrices. Molecularly imprinted polymers (MIPs) have been termed as advanced solid-phase (SPE) materials for the selective extraction and preconcentration of hormones in complex matrices. This review explores the preparation and application of MIPs for the determination of steroid hormones in different sample types.

1. Introduction

Exogenous chemicals with the ability to interfere with the normal function of hormones are known as endocrine disrupting compounds (EDCs) [1]. The effects of EDCs include reduced fertility and elevated chances of cancer [2]. One of the most active and potent group of EDCs in the environment is hormones [3]. Hormones that are naturally found in mammals are called endogenous hormones, while foreign hormones are classified as exogenous hormones and can be either natural or synthetic [4]. Steroids are the most active endocrine hormones found in the environment [1].
Animals produce hormones in endocrine glands, organs, and tissues. These hormones control a vast amount of activities, including essential ones, such as the regulation of cell activity and mood control [5]. Hormones can be broadly classified as steroids and nonsteroids [6]. Structurally, all steroid hormones contain a characteristic arrangement of four cycloalkane rings [7]. Steroids are further classified into mineralocorticoids (e.g., aldosterone), glucocorticoids (e.g., hydrocortisone), androgens (e.g., testosterone (TST)), estrogens (e.g., β-estradiol (E2)), and progesterone (PRO) [5]. Their chemical structures are shown in Figure 1.
Human and animal excretion often contain natural and exogenous variation of hormones [8]. This is due to the fact that while steroids such as estrogens, mineralocorticoids (aldosterone), glucocorticoids, androgens, and progesterones can be secreted in the adrenal cortex [9], there are still exogenous hormones used as contraceptives [10], medicines [11], and growth promoters [12]. As a result, hormones can end up in the environment via wastewater treatment plant effluent discharge into receiving water bodies [13].
Hormonal compounds in the water can pose a health risk, such as reproductive disorders, feminization, masculinization, infertility, and cancer [2]. Water is a good medium for carrying and distributing polar and semipolar compounds [13]. As a result, incomplete removal of hormone pollutants in wastewater treatment plants means effluent discharge into nearby rivers will introduce hormonal pollution into water systems [14]. Table 1 shows the concentration levels of hormones in water and other matrices in different countries. As can be seen, while mostly prevalent in water, hormones can still be detected in soil, food, and even humans. Unusual exposure to hormonal compounds can result in endocrine disruption that can have negative effects, such as abnormal cell growth of hormone-regulated tissues [15]. This can lead to neoplasia, hyperplasia, and even cancer [15].
The analysis and quantification of hormones in water matrices has predominantly been done using chromatography, namely gas chromatography, due to efficient separation and successful identification [16]. However, the biggest limitation of gas chromatography determination is the need for derivatization and conversion [17]. These manipulations can result in loss of analyte [17]. This has driven a surge in liquid chromatography (LC) methods that do not require chemical pretreatment for hormonal analysis and quantification [18]. LC methods often lack the GC specificity in complex matrices and thus require sample preparation in order to ensure accurate quantification [19].
Sample preparation allows for the preconcentration/isolation of ultratrace pollutants [20]. A vast number of sample preparation methods can be distinguished as being either liquid–liquid or solid-phase extraction [20]. Liquid–liquid extraction uses an organic solvent as the extracting phase [21,22]. In contrast, solid-phase extraction, as the name suggests, uses a solid phase (adsorbent material) to extract substances [23]. Solid-phase extraction (SPE) has been one of the most commonly used sample preparation techniques since its development in the 1980s [23]. Its advantages are based on its simplicity, selectivity, high enrichment factors, ease of automation, and use of different adsorbents [24,25]. Molecularly imprinted polymers have been termed as advanced adsorbent materials for SPE [20,24].
Molecular imprinting was first reported by [26] in a bid to generate artificial receptors. The imprinting process is done by polymerizing a functional monomer in the presence of a template molecule [27]. During polymerization, there is formation of a functional monomer–template complex [28]. This results in a 3D polymer network where a template is trapped. The template can then be removed by washing, thereby leaving cavities within the polymer network that are complementary to the size, shape, and molecular interaction of the template [27].
This review aims to investigate the use of molecularly imprinted polymers in the determination of hormones in different matrices, namely water, food, and biological samples. In addition, the most commonly used MIP synthesis methods, such as precipitation polymerization, and bulk and surface imprinting, are briefly discussed.

2. Global Concentration Levels of Hormones

The increasing concentrations of steroid hormone residues in various matrices in the environment is proof that even with the bans that are currently in place, their increased application is still predominant (Table 1).
In general, global studies have shown that these hormones are frequently detected in waste and surface waters [9,29,32,33,34,36,37,38,39,40,41,42]. The detection of steroid hormones in the environment has been reported to differ from country to country (Table 1). In some cases, their concentrations within the same country also differ depending on the regions or provinces. For example, in South Africa, maximum concentrations for estradiol of 7133 ng/L were detected in wastewaters from Gauteng Province [42], while concentrations of up to 2000 ng/L were reported in KwaZulu Natal Province for estrone (E1), 17-β-estradiol (E2), estriol (E3), 17-α-ethinylestradiol (17-α-EE2), androgens, and progestogens (PRO) [9]. Olatunji et al. [45] reported concentrations ranging from 600 to 45,500 ng/L for E2 and E3 in surface water around animal farms in Western Cape. The presence of steroid hormones in various environmental samples around the world are shown in Table 1. Maximum concentrations of up to 14.1 μg/L for E1, E2, E3, 17α-EE2, PRO, and testosterone (TST) were reported in Brazil [30,41], while Beldean-Galea et al. [34] reported steroid hormone concentrations ranging from 2.6 to 47 μg/L in Romanian water matrices. Other significantly high concentrations of steroid hormones were observed in Hungary (0.33–488 μg/L) [37] and Serbia (12–4808 ng/L) [43]. From these studies, it can be noted that wastewater treatment plants (WWTPs) are the main source of pollution of water by steroid hormones. This is because effluent from wastewater plants is often released into nearby rivers. Additionally, WWTPs are not able to completely remove most emerging pollutants, including hormones, during the treatment process [46]. In other instances, such as the study by Olatunji and coworkers [45], farm waste, wash water discharge, and incorrect disposal of unused products were found to be responsible for steroid hormones in surface water around farm lands. Table 2 also shows that steroid hormones have been detected in a variety of sample types. These include surface [39], ground [35], and wastewater [29] as well a food [32], soil [36], and human excreta [44]. This not only increases the chance of pollution but also means that exposure of these potentially endocrine disruptors is increased.

3. Molecularly Imprinted Polymer Synthesis Methods

Molecularly imprinted polymers (MIPs) offer a synthetic route to developing tailorable stationary phases, which are particularly useful in solid-phase extraction [60]. MIPs are synthetic polymers that can rebind a target molecule even in the presence of interferences [61], and they are made by copolymerizing functional monomers in the presence of a template molecule or substitute and a cross-linker [62]. The general preparative method for molecularly imprinted polymers is presented in Figure 2. The functional monomer is chosen based on its affinity for the template, which may be based on chemical properties as well as molecular modelling routines [63]; subsequent polymerization produces a three-dimensional polymer network with binding sites that are ideally complementary to the template in scale, shape, and functionalities [61].
The type of functional monomer has a significant impact on the properties of the MIP being synthesized. The relationship between the functional monomer and the prototype should be either covalent or noncovalent [65]. In 2016, Figueiredo and coworkers identified methylacrylic acid, acrylic acid, and 4 vinylpyridine as the most common functional monomers used in imprinting [65]. The use of methylacrylic acid, acrylic acid, and 4 vinylpyridine, particularly for hormone application, could be due to high favorability of the resulting MIPs [63]. The stability of the functional monomer–template complex, the ability to form hydrogen bonds, and the absence of polymerizable groups are all important factors in MIP synthesis. The crosslinker, which influences MIP morphology, is the final component required for MIP synthesis. It also helps to keep the binding cavities of MIPs stable. The reviews by Spivak and Vasapollo and coworkers [66,67] highlight cross-linkers that are compliant with MIPs.
In general, three separate MIP synthesis approaches have been reported in the literature: covalent, semicovalent, and noncovalent synthesis [61]. The covalent method involves forming prepolymerization complexes in which the template is covalently bound to the monomers and then removing the template through chemical cleavage [68]. The covalent synthesis results in a homogeneous binding site population and limited nonspecific binding sites due to the high stability of the monomer–template interaction [67]. However, reversible cleavage of the complex under mild conditions is difficult to achieve because of covalent monomer–template interactions [69].
The semicovalent approach is an immediate solution to the covalent approach restriction. In this approach, the monomer–template complex rebinding is based on noncovalent interactions, thereby making reversible cleavage simple [40].
Noncovalent imprinting allows the use of weak interactions and is regarded as a more versatile option [69]. The interactions responsible for the self-assembly between the template and monomer include hydrogen bonding, π–π bonding, and electrostatic and hydrophobic interactions [70]. The success of imprinting is characterized by high-affinity binding sites and is said to be dependent on the choice of the functional monomer [71]. Chen and colleagues [71] discussed in detail the different synthesis methods for MIPs. Hence, only precipitation, bulk, and surface imprinted polymerization will be covered in this review.

3.1. Precipitation Polymerization

One preparative phase is needed for precipitation polymerization. This method of polymerization yields uniform and spherical particles (diameters usually less than 1 µm), but it necessitates a significant amount of template [72]. Precipitation polymerization is a surfactant-free process that involves polymerizing monomers in dilute solutions (without overlap or coalescence) and removing the resulting polymer particles from the solution [73]. Entropic precipitation of nanogel (seed) particles, followed by continuous capture of oligomers from solution, is the most common way for particles to develop. The general scheme for precipitation polymerization in shown in Figure 3. In contrast to bulk polymerization, this form of polymerization necessitates a significant amount of solvent [74]. It should also be noted that many variables, such as the polarity of the solvent, reaction temperature, and stirring speed, affect the size of the particles obtained, so reaction conditions should be carefully regulated [75].

3.2. Bulk Imprinting

Bulk imprinting involves imprinting a template molecule in its entirety in a polymer matrix and then fully removing it from the molecularly imprinted substance after polymerization [76]. The bulk polymer is mechanically crushed, and the particles formed are then fractionated in the next step to form small particles from these bulk polymers. For relatively smaller molecules, bulk imprinting is preferred. Due to the ease of adsorption and release of the template molecule, reversible binding can be carried out, which provides the potential for several rounds of reuse [77].
In sensor applications, using a whole polymer as a prototype has certain advantages over other methods [73]. Because the template protein (and the target at the same time) is fully imprinted, the template structure would be very similar to the target structure. However, the approach has certain disadvantages when dealing with larger structures, such as proteins, living cells, and microorganisms. Maintaining the conformational stability of a protein during the polymerization process is difficult [4,78,79]. Furthermore, because of the size of the template, large imprinted sites may be attractive to smaller polypeptides, resulting in cross-reactivity and decreased selectivity [78]. Because of the thick morphology of bulk imprints, large template molecules are embedded too deeply in the matrices, limiting or preventing target molecules from binding to the sites. Low accessibility causes significantly longer response times, drift issues, and poor regeneration A more drastic situation is that removing the target molecule from the MIP is difficult. This will result in hindered binding or, in the worst-case scenario, no binding at all. Alternative imprinting methods, such as surface imprinting, have been designed to address these limitations [79,80].

3.3. Surface Imprinting

Surface imprinting is a useful technique for depositing a thin layer of polymeric material on a variety of substrates, including carbon nanotubes (CNTs), Fe3O4, TiO2, and SiO2. More accessible adsorption sites, rapid mass transfer, fast binding kinetics, and high selectivity are all provided by this layer of MIPs with imprinted cavities on the surface of the particles [72,81]. A general scheme of the synthesis of surface imprinting on a silica nanoparticle core is presented in Figure 4. Although bulk imprinting with these methods is the most popular synthesis technique, more recently, so-called surface imprinting methods have revealed distinct advantages. Such advantages include minimal material waste, enhanced access to binding sites exclusively on the particle’s surface, tailorability of bead size from micro to nanoscale, access to sophisticated core–shell configurations (e.g., inorganic silica core particle with nanothin MIP shell), and reduced mass transfer limitations, leading to rapid binding kinetics [79,81,82].
By establishing binding sites near to and/or at the substrate level, surface imprinting improves the interaction between the template molecule and MIP, ensuring effective mass transfer. For imprinting larger molecules, surface imprinting is an especially promising technique [79]. Surface imprinting also has the advantage of requiring less template during the polymerization process than traditional bulk imprinting. Several imprinting methods have been investigated, including lithographic imprinting [83], dispersed-phase polymerization [84], and grafting via core–shell imprinting [85]. Additionally, electrochemical imprinting is another form of surface imprinting.
In electrochemical polymerization, the solution used contains the template, solvent, functional monomer, and supporting electrolyte [86,87]. There is no use of a core nanoparticle as in traditional surface imprinting polymerization. Electropolymerization is divided into potentiodynamic, potentiostatic, and galvanostatic electropolymerization. The resulting polymers may be neutral or charged due to the movement of solvated counter ion into or out of the film upcharging and discharging during film growth [86]. MIPs prepared using electropolymerization can easily adhere to transducer surfaces, the preparation is rapid, and the film thickness can be controlled [87,88]. This makes them good candidates for sensor applications. The most attractive feature of molecular imprinting using electropolymerization is the complete removal of the template molecule by overoxidation [64,89,90].

4. Sorbent-Based MIP Applications

4.1. Water Samples

Lu and Xu [89] conducted a study investigating the concentrations of E1, 17β-estradiol (E2), and estriol (E3) in tap, river, and lake water samples. In their work, they explored the use of E1-imprinted Fe3O4@SiO2@mSiO2 (MM–MIPs) as an adsorbent DSPE for selective preconcentration and specific recognition of E1, E2, and E3. The MM–MIPs sorbent presented high adsorption capacity, high extraction efficiency, and fast mass transfer for the target analytes. The MM–MIP–SPE combined with HPLC–PDA showed relatively good analytical characteristics. The recoveries ranged from 85 to 95%, and the precision of the method was less than 6%. Low detection limits ranging from 0.09 to 0.4 µg L−1 and high enrichment factor of 1700 were obtained. The use of magnetic MIP was also recently reported by Guc and Schroeder [64], MIPs and magnetic MIPs prepared using bulk polymerization and core–shell method procedures, respectively, were used as adsorbents for selective extraction and preconcentration of E1 and E2 from water samples. The quantification of the analytes in environmental samples was achieved using electrospray ionization mass spectrometry (ESI–MS) and flowing atmospheric pressure after glow mass spectrometry (FAPA–MS). The results obtained revealed that FAPA–MS (LODs = 0.135 µg L−1) was more sensitive that ESI–MS (LODs ranging from 13.6 to 27 µg L−1). The combination of MIP/magnetic MIP–SPE method (methodology scheme in Figure 5) with FAPA–MS resulted in accurate quantification of trace amounts of E1 and E2 in water, and the concentrations were found to be 0.271–0.273 µg L−1. These methods combine the features of MIPs, which ensures the production of selective cavities on the surface of the iron oxide core. This provides magnetic properties to the MIPs, thus facilitating easy removal and reuse. The recognition sites located on the surface of each magnetic MIP particle result in increased selectivity and sensitivity.
In another study, the MIP–SPE method was coupled with high-performance liquid chromatography coupled with diode array detection (HPLC–PDA) for selective determination of E1, E2, E3, 17-α-EE2, PRO, and TST in water [91]. The method proved to be suitable for quantification of steroid hormones in water because it had relatively low LODs in the range of 0.0182–0.0898 μg/mL. In addition, high recoveries (79–101%) and low matrix effects (<20%) suggested that the developed MIP–SPE method could be used for selective identification and quantification of the target analytes [91].
Guedes-Alonso et al. [90] reported the synthesis and application of MIP as an adsorbent for SPE of estrogens from wastewater collected from a veterinary hospital and a wastewater treatment plant. The developed method demonstrated adequate LOD (0.180.45 ng/mL), acceptable recoveries (>60%), and high precision (<RSD 10%). The coupling of MIP–SPE with UPLC enabled accurate quantification of the analytes at concentrations ranging from 1.35 to 2.57 ng/mL.
The consolidated summary of the application of MIPs as adsorbent for the extraction and preconcentration of different types of steroid hormones is presented in Table 2. As can be seen, the use of MIPs as selective adsorbents allows the use of conventional techniques such as HPLC–UV (DAD) or HPLC–FLD for simultaneous determination of hormones in environmental matrices. Furthermore, it can be noted that acceptable to low detection limits (0.04–1.5 µg/L) were obtained using HPLC–DAD/FLD [47,49,50,52,53,54,55,56]. The introduction of MIPs to various analytical techniques has brought much improvement in terms of selectivity and sensitivity. This can be attributed to the properties of MIPs such as the cavities formed, which are complementary to the template’s structural signature (size, shape, and functional group positioning) [64]. The studies conducted by [55,56,58,59,92] examined different steroid hormones, such as E2, beta, and alpha-zearalonol. The application of MIPs as a sorbent in SPE (Table 2) shows that low limits of detection for instruments such as HLPC can be achieved using MIPs.

4.2. Food Samples

Residues of steroid hormones have been detected in various food matrices, especially in milk samples (Table 3). This might be due to the use of steroid hormones as growth promoters. For instance, Tang and colleagues [93] reported the dummy molecularly imprinted polymer microspheres (DMIPMS) as adsorbent for extraction and preconcentration of natural and synthetic estrogens E1, 17β-E2, E3, EE2, DS, DES, and HEX in milk samples. The microspheres were synthesized via Pickering emulsion polymerization, and genistein (GEN) was employed as a dummy template molecule. The FTIR analysis confirmed that the DMIPMS were successfully prepared as all the expected functional groups were observed. The developed method was found to be selective toward the selected seven estrogens, and their quantification was conducted using HPLC–MS/MS. The DMIP–SPE coupled with HPLC–MS/MS displayed excellent linearity with LODs in the range of 0.10–0.35 µg L−1. The recoveries after spiking the milk samples at three levels were between 88.9 and 102.3%. These results suggested that DMIPMS-based SPE could be used for monitoring of trace estrogens in food samples such as milk.
A restricted access media–MIP (RAM–MIP) was reported by Wang and coworkers [93] for selective extraction of 17β-E2 from milk samples. RAM–MIP was prepared via the surface imprinting method whereby monodisperse crosslinked poly(glycidyl methacrylate-co-ethylene glycol dimethacrylate) microspheres were used as the carrier and acryloyl chloride-modified β-cyclodextrin as the hydrophilic functional monomer. The resultant adsorbent was found to have high adsorption affinity toward E2. The RAM–MIP–DSPE coupled with HPLC–PDA was used for analysis of E2 in milk samples. The method showed promising analytical performance, such as high recoveries (up to 95%), high precision (<4%), and relatively high sensitivity (LOD = 2.1 µg/L). In another study, Zhu and colleagues [104] reported the synthesis of zipper-like on/off switchable magnetic molecularly imprinted microspheres (SM–MIMs) as an adsorbent for the analysis of E2 in milk samples (methodology of the scheme is presented in Figure 6). The SM–MIMs was prepared by surface polymerization of acrylamide (AAm) and 2-acrylamide-2-methyl propanesulfonic acid (AMPS). It was observed that the adsorption of E2 to the adsorbent was temperature dependent. The authors reported that this phenomenon was due to the interactions between two polymers (poly(AAm) (PAAm) and poly(AMPS) (PAMPS)), which had on/off switchable property to temperature. The SM–MIM–SPE combined with HPLC–PDA displayed attractive analytical performance, such as high selectivity, low LOD (2.5 µg/L), and high recoveries (79–90%), of E2 in complex matrices. Table 3 also shows that the use of MIPs allows researchers to use simple and inexpensive detection techniques, such as ultraviolet spectrophotometry (UV) [96] and high-performance liquid chromatography with UV detectors [95,97,100,103] and achieve low limits of detection. This demonstrates that the use of MIPs can reduce the dependency on GC analysis.

4.3. Biological and Other Complex Samples

Qiu et al. [105] investigated the levels of anabolic steroids, such as androsterone, stanolone, androstenedione and methyltestosterone, in human urine. This was achieved using MIP-coated SPME fibers combined with GC–MS. In their study, a testosterone MIP was prepared using thermal radical copolymerization of MAA and trimethylolpropanetrimethacrylate (TRIM). The developed method presented promising analytical figures of merit. The LOQs, precision, linearity, and recoveries were in the range of 0.01–0.08 ng/mL, 4–15%, 0.02–1 ng/mL, and 87–108%, respectively. The authors reported enrichment factors, i.e., the enrichment of the chromatographic peaks between these results suggested that the developed MIP-coated SPME/GC–MS was suitable for rapid extraction and determination of trace anabolic steroids in biological matrices.
A selective extraction and determination of PRO hormones from biological matrices such as urine and blood samples were reported by Nezhadali et al. [106]. The extraction and preconcentration of target analytes were achieved using a polypyrrole MIP (prepared via bulk polymerization) as adsorbent followed by GC–FID quantification. Under optimized conditions, the LOD, LOQ, and recoveries were 0.63, 1.9 ng/mL, and 86–101%, respectively. The method was successfully applied for the determination of the target analyte in urine and blood.
Du et al. [107] reported the preparation of dexamethasone-imprinted polymers (DEXA-MIP) via surface molecular imprinting. The synthesis and application summary can be seen in Figure 7. The surface molecular imprinting was achieved using a method called reversible addition–fragmentation chain transfer polymerization on the surface of magnetic nanoparticles. The prepared MIP was used as a magnetic adsorbent for SPE of DEXA from skincare cosmetic samples prior to HPLC–PDA determination. The developed method displayed relatively good accuracy (93–97.6%), high precision (RSD < 3%), and low LODs (0.05 μg/mL). Furthermore, the developed MIP–SPE/HPLC method possessed attractive features, such as specific molecular recognition, high adsorption affinity and selectivity, and simplicity, and it was considered as a good candidate for monitoring trace concentration of DEXA in various complex matrices.
A study by Xu et al. [54] reported the application of dual-template MIP–SBSE combined with HPLC–DAD for analysis of E1, E2, and DES in plastic samples. The developed methods were found to be suitable for selective determination of steroid hormones in complex matrices with recoveries ranging from 78 to 97%. In cases where analytes are different structurally, MIPs are not suitable for analysis of target analytes in complex real samples, such as cosmetics [108]. To overcome these shortcomings, dual-template MIPs are used. For instance, Liu et al. [108] conducted a study investigating the presence of glucocorticoids in cosmetics samples. In their study, they explored the use of novel dual-template magnetic MIP as an adsorbent for SPE. The magnetic MIP was synthesized using the surface polymerization method, and hydrocortisone and DEXA were used as templates. The prepared dual-template magnetic MIP had high affinity toward target analytes, and it was used for extraction and enrichment of hydrocortisone and DEXA in cosmetic products. The magnetic MIP–SPE/HPLC method displayed satisfactory recoveries ranging 86.8–107.5% as well as good precision (RSD <3%). Other applications of MIP based SPE for preconcentration and extraction of hormones prior HPLC analysis are presented in Table 4. As seen in this Table, SPE based MIP enabled accurate quantification of various steroid hormones from urine and serum us-ing less sensitive such as HPLC-UV. In comparison with UHPLC-MS/MS, HPLC-UV had high detection limits (25-92µg/L).
The benefits of using MIPs in solid-phase extraction include improvement of the recognition selectivity, simplicity, flexibility, and detection sensitivity of the extraction process and the solventless nature associated with solid-phase extraction [112,113] MIPs in solid-phase extraction provides an important tool for chemo/bioanalysis in complex matrices and benefits from distinguished advantages, such as easy operation, high throughput, low cost, high selectivity, and durability [114]. However, there are also disadvantages, such as the lack of compatibility between the solvent needed to desorb analytes from the MIP and the mobile phase used (typical drawback of online MISPE protocols) [115].

5. MIP Challenges

Though inexpensive and easy to scale up after calibration of a particular setup, MIP synthesis needs to be highly customized to the desired target(s). This can be a challenging task as there is no universal preparation protocol that ensures adequate selectivity and the MIP technology is not easily transferable among different applications [116,117]. The removal of the template after successful imprinting is important for the steps following it, such as assessing binding capability and nonspecific adsorption on nonimprinted polymers [118]. Because different templates and complementary functional monomers interact differently, techniques for template removal and assessment of binding capability and nonspecific adsorption on nonimprinted polymers are often inconsistent [116]. For example, different polymers necessitate different solvent strength for template removal. Acrylic acid polymers are more resistant than self-polymerizing PDA MIPs [119]. Although their stability is also advantageous in bioapplications because they can be sterilized and reused, the verification of template elimination in is often inconsistent [116]. While some studies have evaluated template removal using separation methods such as HPLC, which is evaluated indirectly as a residual of analyte in the sample that is not bound by MIP, others have not found any precise technique for confirming template removal [117]. Additionally, MIPs do not produce any signal to show analyte binding onto the polymer, so they must be used in conjunction with a suitable detection method [73]. MIP surfaces are often formed over a functional NP, such as one with optical properties. Even though fluorescence and luminescence methods are becoming more common, the problem of determining the optimal penetration depth of light emission for in vivo applications has yet to be solved. When used for whole-body optical imaging, however, the signal may be insufficient [73,120].

6. Conclusions

Growing interest in the use of selective adsorbents for extraction of environmental pollutants, especially those with endocrine-disrupting properties, has resulted in the development of MIP-based SPE methods. The number of publications reporting the use of MIP-based SPE in environmental analysis suggest that MIP-based sorbents remain one of the main factors in the field of sample preparation. This review summarized the recent application of MIP-based methods for selective extraction and preconcentration of steroid hormones as well as sample clean-up of complex matrices, such as soil, food, and biological samples. Previous studies have revealed that the use of MIP-based SPE methods enable accurate quantification of steroid hormones in water, food, biological, and other complex samples. Furthermore, the literature shows that precipitation, bulk, and surface imprinting are the most frequently used methods for the preparation of MIPs. However, the absence of universal synthesis protocols that do not require method tuning and do not have inconsistent template molecule removal after synthesis or nonspecific adsorption on the polymer has become a major challenge associated with the preparation and application of MIPs. Researchers have developed novel hybrid MIP-based sorbents to address these challenges. More recently, the use of sensors is likely to be part of the next generation of analysis methods, especially MIP-based sensors. This is because the synthesis methods used for the preparation of MIP sensors, such as electropolymerization, also addresses one of the most important challenges of MIP synthesis, which is related to the removal of the template. Finally, the use of sensors would reduce costs associated with analysis as they eliminate the need for sophisticated instrumentation. The use of MIPs as selective adsorbents for steroid hormone detection could still be the driver of the next set of innovations.

Author Contributions

Conceptualization, A.M., B.M. and P.N.N.; methodology, A.M. and P.N.N.; investigation, A.M., S.K.S., B.M. and P.N.N.; resources, B.M. and P.N.N.; writing—original draft preparation, A.M. and S.K.S.; writing—review and editing, A.M., S.K.S., B.M. and P.N.N.; visualization, A.M., S.K.S. and P.N.N.; supervision, B.M. and P.N.N.; project administration, B.M. and P.N.N.; funding acquisition, A.M., B.M. and P.N.N. All authors have read and agreed to the published version of the manuscript.

Funding

This research was funded by the National Research Foundation (NRF, South Africa, grant nos. 113010 and 91230).

Informed Consent Statement

Not applicable.

Acknowledgments

The authors acknowledge University of Johannesburg (Department of Chemical Sciences) and Ulm University (Institute of Analytical and Bioanalytical Chemistry).

Conflicts of Interest

The authors declare no conflict of interest. The funders had no role in the design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript; or in the decision to publish the results.

References

  1. Aufartová, J.; Mahugo-santana, C.; Sosa-ferrera, Z.; Santana-rodríguez, J.J.; Nováková, L.; Solich, P. Determination of steroid hormones in biological and environmental samples using green microextraction techniques: An overview. Anal. Chim. Acta 2011, 704, 33–46. [Google Scholar] [CrossRef] [PubMed]
  2. Casals-Casas, C.; Desvergne, B. Endocrine disruptors: From endocrine to metabolic disruption. Annu. Rev. Physiol. 2011, 73, 135–162. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  3. Pedrouzo, M.; Borrull, F.; Pocurull, E.; Marcé, R.M. Presence of pharmaceuticals and hormones in waters from sewage treatment plants. Water Air Soil Pollut. 2011, 217, 267–281. [Google Scholar] [CrossRef]
  4. Avar, P.; Maász, G.; Takács, P.; Lovas, S.; Zrínyi, Z.; Svigruha, R.; Takátsy, A.; Tóth, L.G.; Pirger, Z. HPLC-MS/MS analysis of steroid hormones in environmental water samples. Drug Test. Anal. 2016, 8, 123–127. [Google Scholar] [CrossRef] [PubMed]
  5. Nezami, A.; Nosrati, R.; Golichenari, B.; Rezaee, R. Nanomaterial-based aptasensors and bioaf fi nity sensors for quantitative detection of 17 b -estradiol. Trends Anal. Chem. 2017, 94, 95–105. [Google Scholar] [CrossRef]
  6. Di Donna, L.; Benabdelkamel, H.; Taverna, D.; Indelicato, S.; Aiello, D.; Napoli, A.; Sindona, G.; Mazzotti, F. Determination of ketosteroid hormones in meat by liquid chromatography tandem mass spectrometry and derivatization chemistry. Anal. Bioanal. Chem. 2015, 407, 5835–5842. [Google Scholar] [CrossRef]
  7. Puckowski, A.; Mioduszewska, K.; Łukaszewicz, P.; Borecka, M.; Caban, M.; Maszkowska, J.; Stepnowski, P. Bioaccumulation and analytics of pharmaceutical residues in the environment: A review. J. Pharm. Biomed. Anal. 2016, 127, 232–255. [Google Scholar] [CrossRef]
  8. Fang, T.Y.; Praveena, S.M.; deBurbure, C.; Aris, A.Z.; Ismail, S.N.S.; Rasdi, I. Analytical techniques for steroid estrogens in water samples-A review. Chemosphere 2016, 165, 358–368. [Google Scholar] [CrossRef] [PubMed]
  9. Manickum, T.; John, W. The current preference for the immuno-analytical ELISA method for quantitation of steroid hormones (endocrine disruptor compounds) in wastewater in South Africa. Anal. Bioanal. Chem. 2015, 407, 4949–4970. [Google Scholar] [CrossRef] [PubMed]
  10. Zhang, Z.; Feng, Y.; Gao, P.; Wang, C.; Ren, N. Occurrence and removal efficiencies of eight EDCs and estrogenicity in a STP. J. Environ. Monit. 2011, 13, 1366–1373. [Google Scholar] [CrossRef] [PubMed]
  11. Ye, X.; Guo, X.; Cui, X.; Zhang, X.; Zhang, H.; Wang, M.K.; Qiu, L.; Chen, S. Occurrence and removal of endocrine-disrupting chemicals in wastewater treatment plants in the Three Gorges Reservoir area, Chongqing, China. J. Environ. Monit. 2012, 14, 2204–2211. [Google Scholar] [CrossRef] [PubMed]
  12. Fernández-Arauzo, L.; Pimentel-Trapero, D.; Hernández-Carrasquilla, M. Simultaneous determination of resorcylic acid lactones, β and α trenbolone and stilbenes in bovine urine by UHPLC/MS/MS. J. Chromatogr. B Anal. Technol. Biomed. Life Sci. 2014, 973, 89–96. [Google Scholar] [CrossRef]
  13. Białk-Bielińska, A.; Kumirska, J.; Borecka, M.; Caban, M.; Paszkiewicz, M.; Pazdro, K.; Stepnowski, P. Journal of Pharmaceutical and Biomedical Analysis Selected analytical challenges in the determination of pharmaceuticals in drinking/marine waters and soil/sediment. J. Pharm. Biomed. Anal. 2016, 121, 271–296. [Google Scholar] [CrossRef] [PubMed]
  14. Matić, I.; Grujić, S.; Jauković, Z.; Laušević, M. Trace analysis of selected hormones and sterols in river sediments by liquid chromatography-atmospheric pressure chemical ionization–tandem mass spectrometry. J. Chromatogr. A 2014, 1364, 117–127. [Google Scholar] [CrossRef]
  15. Hamid, H.; Eskicioglu, C. Fate of estrogenic hormones in wastewater and sludge treatment: A review of properties and analytical detection techniques in sludge matrix. Water Res. 2012, 46, 5813–5833. [Google Scholar] [CrossRef] [PubMed]
  16. Pérez, R.L.; Escandar, G.M. Multivariate calibration-assisted high-performance liquid chromatography with dual UV and fluorimetric detection for the analysis of natural and synthetic sex hormones in environmental waters and sediments. Environ. Pollut. 2016, 209, 114–122. [Google Scholar] [CrossRef] [PubMed]
  17. Naldi, A.C.; Fayad, P.B.; Prévost, M.; Sauvé, S. Analysis of steroid hormones and their conjugated forms in water and urine by on—line solid—phase extraction coupled to liquid chromatography tandem mass spectrometry. Chem. Cent. J. 2016, 1–17. [Google Scholar] [CrossRef] [Green Version]
  18. Tomšíková, H.; Aufartová, J.; Solich, P.; Nováková, L.; Sosa-Ferrera, Z.; Santana-Rodríguez, J.J. High-sensitivity analysis of female-steroid hormones in environmental samples. TrAC Trends Anal. Chem. 2012, 34, 35–58. [Google Scholar] [CrossRef]
  19. Migowska, N.; Caban, M.; Stepnowski, P.; Kumirska, J. Simultaneous analysis of non-steroidal anti-inflammatory drugs and estrogenic hormones in water and wastewater samples using gas chromatography–mass spectrometry and gas chromatography with electron capture detection. Sci. Total Environ. 2012, 441, 77–88. [Google Scholar] [CrossRef]
  20. Buszewski, B.; Szultka, M. Past, present, and future of solid phase extraction: A review. Crit. Rev. Anal. Chem. 2012, 42, 198–213. [Google Scholar] [CrossRef]
  21. Matamoros, V.; Calderón-Preciado, D.; Domínguez, C.; Bayona, J.M. Analytical procedures for the determination of emerging organic contaminants in plant material: A review. Anal. Chim. Acta 2012, 722, 8–20. [Google Scholar] [CrossRef]
  22. Buchberger, W.W. Current approaches to trace analysis of pharmaceuticals and personal care products in the environment. J. Chromatogr. A 2011, 1218, 603–618. [Google Scholar] [CrossRef]
  23. Dimpe, K.M.; Nomngongo, P.N. Current sample preparation methodologies for analysis of emerging pollutants in different environmental matrices. TrAC Trends Anal. Chem. 2016, 82, 199–207. [Google Scholar] [CrossRef]
  24. Żwir-Ferenc, A.; Biziuk, M. Solid Phase Extraction Technique--Trends, Opportunities and Applications. Polish J. Environ. Stud. 2006, 15, 677–690. [Google Scholar]
  25. Sosa-Ferrera, Z.; Mahugo-Santana, C.; Santana-Rodríguez, J.J. Analytical Methodologies for the Determination of Endocrine Disrupting Compounds in Biological and Environmental Samples. Biomed Res. Int. 2013, 2013, 1–23. [Google Scholar] [CrossRef]
  26. Wulf, G.; Sarhan, A. Macromolecular colloquim, Angevv. Chem. Int. Ed. Engl. 1972, 11, 341. [Google Scholar]
  27. Wackerlig, J.; Schirhagl, R. Applications of molecularly imprinted polymer nanoparticles and their advances toward industrial use: A review. Anal. Chem. 2016, 88, 250–261. [Google Scholar] [CrossRef]
  28. Cheong, W.J.; Yang, S.H.; Ali, F. Molecular imprinted polymers for separation science: A review of reviews. J. Sep. Sci. 2013, 36, 609–628. [Google Scholar] [CrossRef] [PubMed]
  29. Fang, T.Y.; Praveena, S.M.; Aris, A.Z.; Ismail, S.N.S.; Rasdi, I. Quantification of selected steroid hormones (17β-Estradiol and 17α-Ethynylestradiol) in wastewater treatment plants in Klang Valley (Malaysia). Chemosphere 2019, 215, 153–162. [Google Scholar] [CrossRef]
  30. Moraes, F.C.; Rossi, B.; Donatoni, M.C.; de Oliveira, K.T.; Pereira, E.C. Sensitive determination of 17β-estradiol in river water using a graphene based electrochemical sensor. Anal. Chim. Acta 2015, 881, 37–43. [Google Scholar] [CrossRef]
  31. Guo, F.; Liu, Q.; Qu, G.; Song, S.; Sun, J.; Shi, J.; Jiang, G. Simultaneous determination of five estrogens and four androgens in water samples by online solid-phase extraction coupled with high-performance liquid chromatography–tandem mass spectrometry. J. Chromatogr. A 2013, 1281, 9–18. [Google Scholar] [CrossRef]
  32. Su, R.; Wang, X.; Xu, X.; Wang, Z.; Li, D.; Zhao, X.; Li, X.; Zhang, H.; Yu, A. Application of multiwall carbon nanotubes-based matrix solid phase dispersion extraction for determination of hormones in butter by gas chromatography mass spectrometry. J. Chromatogr. A 2011, 1218, 5047–5054. [Google Scholar] [CrossRef]
  33. Silva, C.P.; Lima, D.L.D.; Schneider, R.J.; Otero, M.; Esteves, V.I. Development of ELISA methodologies for the direct determination of 17β-estradiol and 17α-ethinylestradiol in complex aqueous matrices. J. Environ. Manag. 2013, 124, 121–127. [Google Scholar] [CrossRef] [PubMed]
  34. Beldean-Galea, M.S.; Klein, R.; Coman, M.-V. Simultaneous Determination of Four Nonsteroidal Anti-Inflammatory Drugs and Three Estrogen Steroid Hormones in Wastewater Samples by Dispersive Liquid–Liquid Microextraction Based on Solidification of Floating Organic Droplet and HPLC. J. AOAC Int. 2020, 103, 392–398. [Google Scholar] [CrossRef] [PubMed]
  35. Kotowska, U.; Kapelewska, J.; Kotowski, A.; Pietuszewska, E. Rapid and sensitive analysis of hormones and other emerging contaminants in groundwater using ultrasound-assisted emulsification microextraction with solidification of floating organic droplet followed by GC-MS Detection. Water 2019, 11, 1638. [Google Scholar] [CrossRef] [Green Version]
  36. Zhang, F.-S.; Xie, Y.-F.; Li, X.-W.; Wang, D.-Y.; Yang, L.-S.; Nie, Z.-Q. Accumulation of steroid hormones in soil and its adjacent aquatic environment from a typical intensive vegetable cultivation of North China. Sci. Total Environ. 2015, 538, 423–430. [Google Scholar] [CrossRef] [PubMed]
  37. Andrási, N.; Molnár, B.; Dobos, B.; Vasanits-Zsigrai, A.; Záray, G.; Molnár-Perl, I. Determination of steroids in the dissolved and in the suspended phases of wastewater and Danube River samples by gas chromatography, tandem mass spectrometry. Talanta 2013, 115, 367–373. [Google Scholar] [CrossRef] [PubMed]
  38. Zhao, Y.-G.; Zhang, Y.; Zhan, P.-P.; Chen, X.-H.; Pan, S.-D.; Jin, M.-C. Fast determination of 24 steroid hormones in river water using magnetic dispersive solid phase extraction followed by liquid chromatography–tandem mass spectrometry. Environ. Sci. Pollut. Res. 2016, 23, 1529–1539. [Google Scholar] [CrossRef] [PubMed]
  39. Wang, H.-X.; Zhou, Y.; Jiang, Q.-W. Simultaneous screening of estrogens, progestogens, and phenols and their metabolites in potable water and river water by ultra-performance liquid chromatography coupled with quadrupole time-of-flight mass spectrometry. Microchem. J. 2012, 100, 83–94. [Google Scholar] [CrossRef]
  40. Goyon, A.; Cai, J.Z.; Kraehenbuehl, K.; Hartmann, C.; Shao, B.; Mottier, P. Determination of steroid hormones in bovine milk by LC-MS/MS and their levels in Swiss Holstein cow milk. Food Addit. Contam. Part A 2016, 33, 804–816. [Google Scholar] [CrossRef] [PubMed]
  41. MFM Sampaio, N.; DB Castilhos, N.; C da Silva, B.; C Riegel-Vidotti, I.; JG Silva, B. Evaluation of Polyvinyl Alcohol/Pectin-Based Hydrogel Disks as Extraction Phase for Determination of Steroidal Hormones in Aqueous Samples by GC-MS/MS. Molecules 2019, 24, 40. [Google Scholar] [CrossRef] [Green Version]
  42. Mhuka, V.; Dube, S.; Nindi, M.M. Occurrence of pharmaceutical and personal care products (PPCPs) in wastewater and receiving waters in South Africa using LC-OrbitrapTM MS. Emerg. Contam. 2020, 6, 250–258. [Google Scholar] [CrossRef]
  43. Jauković, Z.D.; Grujić, S.D.; Bujagić, I.V.M.; Laušević, M.D. Determination of sterols and steroid hormones in surface water and wastewater using liquid chromatography-atmospheric pressure chemical ionization-mass spectrometry. Microchem. J. 2017, 135, 39–47. [Google Scholar] [CrossRef]
  44. Arismendi, D.; Díaz, K.; Aguilera-Marabolí, N.; Sepúlveda, B.; Richter, P. Rotating-disk sorptive extraction for the determination of sex hormones and triclosan in urine by gas chromatography-mass spectrometry: Clean-up integrated steps and improved derivatization. Microchem. J. 2020, 105149. [Google Scholar] [CrossRef]
  45. Olatunji, O.S.; Fatoki, O.S.; Opeolu, B.O.; Ximba, B.J.; Chitongo, R. Determination of selected steroid hormones in some surface water around animal farms in Cape Town using HPLC-DAD. Environ. Monit. Assess. 2017, 189, 363. [Google Scholar] [CrossRef]
  46. Tang, Z.; Liu, Z.; Wang, H.; Dang, Z.; Liu, Y. Occurrence and removal of 17α-ethynylestradiol (EE2) in municipal wastewater treatment plants: Current status and challenges. Chemosphere 2021, 129551. [Google Scholar] [CrossRef] [PubMed]
  47. Hao, Y.; Gao, R.; Shi, L.; Liu, D.; Tang, Y.; Guo, Z. Water-compatible magnetic imprinted nanoparticles served as solid-phase extraction sorbents for selective determination of trace 17beta-estradiol in environmental water samples by liquid chromatography. J. Chromatogr. A 2015, 1396, 7–16. [Google Scholar] [CrossRef]
  48. Kopperi, M.; Riekkola, M.-L. Non-targeted evaluation of selectivity of water-compatible class selective adsorbents for the analysis of steroids in wastewater. Anal. Chim. Acta 2016, 920, 47–53. [Google Scholar] [CrossRef]
  49. Gao, R.; Su, X.; He, X.; Chen, L.; Zhang, Y. Preparation and characterisation of core–shell CNTs@ MIPs nanocomposites and selective removal of estrone from water samples. Talanta 2011, 83, 757–764. [Google Scholar] [CrossRef] [PubMed]
  50. Yagishita, M.; Kubo, T.; Nakano, T.; Shiraishi, F.; Tanigawa, T.; Naito, T.; Sano, T.; Nakayama, S.F.; Nakajima, D.; Otsuka, K. Efficient extraction of estrogen receptor–active compounds from environmental surface water via a receptor-mimic adsorbent, a hydrophilic PEG-based molecularly imprinted polymer. Chemosphere 2019, 217, 204–212. [Google Scholar] [CrossRef] [PubMed]
  51. Lin, Z.; He, Q.; Wang, L.; Wang, X.; Dong, Q.; Huang, C. Preparation of magnetic multi-functional molecularly imprinted polymer beads for determining environmental estrogens in water samples. J. Hazard. Mater. 2013, 252, 57–63. [Google Scholar] [CrossRef] [PubMed]
  52. Qiao, L.; Gan, N.; Hu, F.; Wang, D.; Lan, H.; Li, T.; Wang, H. Magnetic nanospheres with a molecularly imprinted shell for the preconcentration of diethylstilbestrol. Microchim. Acta 2014, 181, 1341–1351. [Google Scholar] [CrossRef]
  53. Lucci, P.; Núñez, O.; Galceran, M.T. Solid-phase extraction using molecularly imprinted polymer for selective extraction of natural and synthetic estrogens from aqueous samples. J. Chromatogr. A 2011, 1218, 4828–4833. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Xu, Z.; Yang, Z.; Liu, Z. Development of dual-templates molecularly imprinted stir bar sorptive extraction and its application for the analysis of environmental estrogens in water and plastic samples. J. Chromatogr. A 2014, 1358, 52–59. [Google Scholar] [CrossRef] [PubMed]
  55. González, A.; Cerdà, V. Development of an automatic sequential injection analysis-lab on valve system exploiting molecularly imprinted polymers coupled with high performance liquid chromatography for the determination of estrogens in wastewater samples. Talanta 2020, 209, 120564. [Google Scholar] [CrossRef]
  56. González-Sálamo, J.; Socas-Rodríguez, B.; Hernández-Borges, J.; del Mar Afonso, M.; Rodríguez-Delgado, M.Á. Evaluation of two molecularly imprinted polymers for the solid-phase extraction of natural, synthetic and mycoestrogens from environmental water samples before liquid chromatography with mass spectrometry. J. Sep. Sci. 2015, 38, 2692–2699. [Google Scholar] [CrossRef] [PubMed]
  57. Sadowski, R.; Gadzała-Kopciuch, R. Isolation and determination of estrogens in water samples by solid-phase extraction using molecularly imprinted polymers and HPLC. J. Sep. Sci. 2013, 36, 2299–2305. [Google Scholar] [CrossRef] [PubMed]
  58. Chen, W.; Xue, M.; Xue, F.; Mu, X.; Xu, Z.; Meng, Z.; Zhu, G.; Shea, K.J. Molecularly imprinted hollow spheres for the solid phase extraction of estrogens. Talanta 2015, 140, 68–72. [Google Scholar] [CrossRef] [PubMed]
  59. He, X.; Mei, X.; Wang, J.; Lian, Z.; Tan, L.; Wu, W. Determination of diethylstilbestrol in seawater by molecularly imprinted solid-phase extraction coupled with high-performance liquid chromatography. Mar. Pollut. Bull. 2016, 102, 142–147. [Google Scholar] [CrossRef] [PubMed]
  60. Whitcombe, M.J.; Nicholls, I.A. Molecular Imprinting Science and Technology: A Survey of the Literature for the Years from 2004 to 2011. J. Mol. Recognit. 2011, 27, 297–401. [Google Scholar]
  61. Beyazit, S.; Tse, B.; Bui, S.; Haupt, K.; Gonzato, C. Progress in Polymer Science Molecularly imprinted polymer nanomaterials and nanocomposites by controlled / living radical polymerization. Prog. Polym. Sci. 2016, 62, 1–21. [Google Scholar] [CrossRef]
  62. Speltini, A.; Scalabrini, A.; Maraschi, F.; Sturini, M.; Profumo, A. Analytica Chimica Acta Newest applications of molecularly imprinted polymers for extraction of contaminants from environmental and food matrices: A review. Anal. Chim. Acta 2017, 974, 1–26. [Google Scholar] [CrossRef]
  63. Zink, S.; Moura, F.A.; da Silva Autreto, P.A.; Galvao, D.S.; Mizaikoff, B. Efficient prediction of suitable functional monomers for molecular imprinting via local density of states calculations. Phys. Chem. Chem. Phys. 2018, 20, 13153–13158. [Google Scholar] [CrossRef] [PubMed]
  64. Guć, M.; Schroeder, G. Molecularly Imprinted Polymers and Magnetic Molecularly Imprinted Polymers for Selective Determination of Estrogens in Water by ESI-MS/FAPA-MS. Biomolecules 2020, 10, 672. [Google Scholar] [CrossRef]
  65. Figueiredo, L.; Erny, G.L.; Santos, L.; Alves, A. Applications of molecularly imprinted polymers to the analysis and removal of personal care products: A review. Talanta 2016, 146, 754–765. [Google Scholar] [CrossRef] [Green Version]
  66. Spivak, D.A. Optimization, evaluation, and characterization of molecularly imprinted polymers. Adv. Drug Deliv. Rev. 2005, 57, 1779–1794. [Google Scholar] [CrossRef]
  67. Vasapollo, G.; Sole, R.D.; Mergola, L.; Lazzoi, M.R.; Scardino, A.; Scorrano, S.; Mele, G. Molecularly imprinted polymers: Present and future prospective. Int. J. Mol. Sci. 2011, 12, 5908–5945. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  68. Wei, S.; Molinelli, A.; Mizaikoff, B. Molecularly imprinted micro and nanospheres for the selective recognition of 17 β-estradiol. Biosens. Bioelectron. 2006, 21, 1943–1951. [Google Scholar] [CrossRef]
  69. Chen, L.; Li, J. Recent advances in molecular imprinting technology: Current status, challenges and highlighted applications. Chem. Soc. Rev. 2011, 40, 2922–2942. [Google Scholar] [CrossRef]
  70. Caro, E.; Masqué, N.; Marcé, R.M.; Borrull, F.; Cormack, P.A.G.; Sherrington, D.C. Non-covalent and semi-covalent molecularly imprinted polymers for selective on-line solid-phase extraction of 4-nitrophenol from water samples. J. Chromatogr. A 2002, 963, 169–178. [Google Scholar] [CrossRef]
  71. Chen, L.; Wang, X.; Lu, W.; Wu, X.; Li, J. Molecular imprinting: Perspectives and applications. Chem. Soc. Rev. 2016, 45, 2137–2211. [Google Scholar] [CrossRef] [PubMed]
  72. Azizi, A.; Bottaro, C.S. A critical review of molecularly imprinted polymers for the analysis of organic pollutants in environmental water samples. J. Chromatogr. A 2020, 1614, 460603. [Google Scholar] [CrossRef] [PubMed]
  73. Włoch, M.; Datta, J. Synthesis and polymerisation techniques of molecularly imprinted polymers. Compr. Anal. Chem. 2019, 86, 17–40. [Google Scholar]
  74. Phungpanya, C.; Chaipuang, A.; Machan, T.; Watla-iad, K.; Thongpoon, C.; Suwantong, O. Synthesis of prednisolone molecularly imprinted polymer nanoparticles by precipitation polymerization. Polym. Adv. Technol. 2018, 29, 3075–3084. [Google Scholar] [CrossRef]
  75. Beltran, A.; Marcé, R.M.; Cormack, P.A.G.; Borrull, F. Synthesis by precipitation polymerisation of molecularly imprinted polymer microspheres for the selective extraction of carbamazepine and oxcarbazepine from human urine. J. Chromatogr. A 2009, 1216, 2248–2253. [Google Scholar] [CrossRef] [PubMed]
  76. Yang, K.; Li, S.; Liu, L.; Chen, Y.; Zhou, W.; Pei, J.; Liang, Z.; Zhang, L.; Zhang, Y. Epitope imprinting technology: Progress, applications, and perspectives toward artificial antibodies. Adv. Mater. 2019, 31, 1902048. [Google Scholar] [CrossRef] [PubMed]
  77. Zhang, Z.; Liu, J. Molecular Imprinting with Functional DNA. Small 2019, 15, 1805246. [Google Scholar] [CrossRef] [PubMed]
  78. Ma, Y.; Pan, G.; Zhang, Y.; Guo, X.; Zhang, H. Comparative study of the molecularly imprinted polymers prepared by reversible addition–fragmentation chain transfer “bulk” polymerization and traditional radical “bulk” polymerization. J. Mol. Recognit. 2013, 26, 240–251. [Google Scholar] [CrossRef] [PubMed]
  79. Riedel, D.; Mizaikoff, B. Surface Imprinted Micro- and Nanoparticles. In Comprehensive Analytical Chemistry; Marć, M., Ed.; Elsevier: Amsterdam, The Netherlands, 2019; pp. 153–191. [Google Scholar]
  80. Dinc, M.; Esen, C.; Mizaikoff, B. Recent advances on core–shell magnetic molecularly imprinted polymers for biomacromolecules. Trends Anal. Chem. 2019, 14, 202–217. [Google Scholar] [CrossRef]
  81. Zhu, H.; Yao, H.; Xia, K.; Liu, J.; Yin, X.; Zhang, W.; Pan, J. Magnetic nanoparticles combining teamed boronate affinity and surface imprinting for efficient selective recognition of glycoproteins under physiological pH. Chem. Eng. J. 2018, 346, 317–328. [Google Scholar] [CrossRef]
  82. Meier, F.; Schott, B.; Riedel, D.; Mizaikoff, B. Computational and experimental study on the influence of the porogen on the selectivity of 4-nitrophenol molecularly imprinted polymers. Anal. Chim. Acta 2012, 744, 68–74. [Google Scholar] [CrossRef]
  83. Vozzi, G.; Morelli, I.; Vozzi, F.; Andreoni, C.; Salsedo, E.; Morachioli, A.; Giusti, P.; Ciardelli, G. SOFT-MI: A novel microfabrication technique integrating soft-lithography and molecular imprinting for tissue engineering applications. Biotechnol. Bioeng. 2010, 106, 804–817. [Google Scholar] [CrossRef]
  84. Torres, J.J.; Montejano, H.A.; Chesta, C.A. Characterization of imprinted microbeads synthesized via minisuspension polymerization. Macromol. Mater. Eng. 2012, 297, 342–352. [Google Scholar] [CrossRef]
  85. Song, Z.; Meng, M.; Pan, J.; Li, C.; Wang, J.; Dai, J.; Yan, Y. Surface molecularly imprinted polymers based on yeast prepared by atom transfer radical emulsion polymerization for selective recognition of ciprofloxacin from aqueous medium. J. Appl. Polym. Sci. 2013, 131. [Google Scholar] [CrossRef]
  86. Sharma, P.S.; Pietrzyk-Le, A.; D’souza, F.; Kutner, W. Electrochemically synthesized polymers in molecular imprinting for chemical sensing. Anal. Bioanal. Chem. 2012, 402, 3177–3204. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  87. Gonçalves, L.M. Electropolymerized molecularly imprinted polymers (e-MIPs), perceptions based in recent literature for soon-to-be world-class scientists. Curr. Opin. Electrochem. 2020. [Google Scholar] [CrossRef]
  88. Peng, Y.; Su, H. Recent innovations of molecularly imprinted electrochemical sensors based on electropolymerization technique. Curr. Anal. Chem. 2015, 11, 307–317. [Google Scholar] [CrossRef]
  89. Lu, H.; Xu, S. Mesoporous structured estrone imprinted Fe3O4@ SiO2@ mSiO2 for highly sensitive and selective detection of estrogens from water samples by HPLC. Talanta 2015, 144, 303–311. [Google Scholar] [CrossRef] [PubMed]
  90. Guedes-Alonso, R.; Santana-Viera, S.; Sosa-Ferrera, Z.; Santana-Rodríguez, J.J. Molecularly imprinted solid-phase extraction coupled with ultra high performance liquid chromatography and fluorescence detection for the determination of estrogens and their metabolites in wastewater. J. Sep. Sci. 2015, 38, 3961–3968. [Google Scholar] [CrossRef] [PubMed]
  91. Czarny, K.; Szczukocki, D.; Krawczyk, B.; Juszczak, R.; Skrzypek, S.; Gadzała-Kopciuch, R. Molecularly imprinted polymer film grafted from porous silica for efficient enrichment of steroid hormones in water samples. J. Sep. Sci. 2019, 42, 2858–2866. [Google Scholar] [CrossRef] [PubMed]
  92. Hu, Y.; Wang, Y.; Chen, X.; Hu, Y.; Li, G. A novel molecularly imprinted solid-phase microextraction fiber coupled with high performance liquid chromatography for analysis of trace estrogens in fishery samples. Talanta 2010, 80, 2099–2105. [Google Scholar] [CrossRef]
  93. Tang, J.; Wang, J.; Yuan, L.; Xiao, Y.; Wang, X.; Yang, Z. Trace analysis of estrogens in milk samples by molecularly imprinted solid phase extraction with genistein as a dummy template molecule and high-performance liquid chromatography–tandem mass spectrometry. Steroids 2019, 145, 23–31. [Google Scholar] [CrossRef]
  94. Xiong, H.; Wu, X.; Lu, W.; Fu, J.; Peng, H.; Li, J.; Wang, X.; Xiong, H.; Chen, L. Switchable zipper-like thermoresponsive molecularly imprinted polymers for selective recognition and extraction of estradiol. Talanta 2018, 176, 187–194. [Google Scholar] [CrossRef] [PubMed]
  95. Mirzajani, R.; Kardani, F.; Ramezani, Z. A nanocomposite consisting of graphene oxide, zeolite imidazolate framework 8, and a molecularly imprinted polymer for (multiple) fiber solid phase microextraction of sterol and steroid hormones prior to their quantitation by HPLC. Microchim. Acta 2019, 186, 129. [Google Scholar] [CrossRef]
  96. Ning, F.; Peng, H.; Li, J.; Chen, L.; Xiong, H. Molecularly imprinted polymer on magnetic graphene oxide for fast and selective extraction of 17β-estradiol. J. Agric. Food Chem. 2014, 62, 7436–7443. [Google Scholar] [CrossRef]
  97. Gao, G.; Xing, Y.; Liu, T.; Wang, J.; Hou, X. UiO-66(Zr) as sorbent for porous membrane protected micro-solid-phase extraction androgens and progestogens in environmental water samples coupled with LC-MS/MS analysis: The application of experimental and molecular simulation method. Microchem. J. 2019, 146, 126–133. [Google Scholar] [CrossRef]
  98. Gong, Y.; Niu, Y.; Gong, X.; Ma, M.; Ren, X.; Zhu, W.; Luo, R.; Gong, B. Preparation of 17β-estradiol-imprinted material by surface-initiated atom transfer radical polymerization and its application. J. Sep. Sci. 2015, 38, 1254–1261. [Google Scholar] [CrossRef] [PubMed]
  99. Zhang, J.; Ni, Y.; Wang, L.; Ma, J.; Zhang, Z. Selective solid-phase extraction of artificial chemicals from milk samples using multiple-template surface molecularly imprinted polymers. Biomed. Chromatogr. 2015, 29, 1267–1273. [Google Scholar] [CrossRef]
  100. Lan, H.; Gan, N.; Pan, D.; Hu, F.; Li, T.; Long, N.; Qiao, L. An automated solid-phase microextraction method based on magnetic molecularly imprinted polymer as fiber coating for detection of trace estrogens in milk powder. J. Chromatogr. A 2014, 1331, 10–18. [Google Scholar] [CrossRef] [PubMed]
  101. Yuan, L.; Ma, J.; Ding, M.; Wang, S.; Wu, X.; Li, Y.; Ma, K.; Zhou, X.; Li, F. Preparation of estriol–molecularly imprinted silica nanoparticles for determining oestrogens in milk tablets. Food Chem. 2012, 131, 1063–1068. [Google Scholar] [CrossRef]
  102. Qiao, L.; Gan, N.; Wang, J.; Gao, H.; Hu, F.; Wang, H.; Li, T. Novel molecularly imprinted stir bar sorptive extraction based on an 8-electrode array for preconcentration of trace exogenous estrogens in meat. Anal. Chim. Acta 2015, 853, 342–350. [Google Scholar] [CrossRef]
  103. Wang, L.; Yan, H.; Yang, C.; Li, Z.; Qiao, F. Synthesis of mimic molecularly imprinted ordered mesoporous silica adsorbent by thermally reversible semicovalent approach for pipette-tip solid-phase extraction-liquid chromatography fluorescence determination of estradiol in milk. J. Chromatogr. A 2016, 1456, 58–67. [Google Scholar] [CrossRef]
  104. Zhu, W.; Peng, H.; Luo, M.; Yu, N.; Xiong, H.; Wang, R.; Li, Y. Zipper-like magnetic molecularly imprinted microspheres for on/off-switchable recognition and extraction of 17β-estradiol from food samples. Food Chem. 2018, 261, 87–95. [Google Scholar] [CrossRef]
  105. Qiu, L.; Liu, W.; Huang, M.; Zhang, L. Preparation and application of solid-phase microextraction fiber based on molecularly imprinted polymer for determination of anabolic steroids in complicated samples. J. Chromatogr. A 2010, 1217, 7461–7470. [Google Scholar] [CrossRef] [PubMed]
  106. Nezhadali, A.; Es’haghi, Z.; Khatibi, A. Selective extraction of progesterone hormones from environmental and biological samples using a polypyrrole molecularly imprinted polymer and determination by gas chromatography. Anal. Methods 2016, 8, 1813–1827. [Google Scholar] [CrossRef]
  107. Du, W.; Zhang, B.; Guo, P.; Chen, G.; Chang, C.; Fu, Q. Facile preparation of magnetic molecularly imprinted polymers for the selective extraction and determination of dexamethasone in skincare cosmetics using HPLC. J. Sep. Sci. 2018, 41, 2441–2452. [Google Scholar] [CrossRef] [PubMed]
  108. Liu, M.; Li, X.; Li, J.; Wu, Z.; Wang, F.; Liu, L.; Tan, X.; Lei, F. Selective separation and determination of glucocorticoids in cosmetics using dual-template magnetic molecularly imprinted polymers and HPLC. J. Colloid Interface Sci. 2017, 504, 124–133. [Google Scholar] [CrossRef]
  109. de Oliveira, H.L.; Pires, B.C.; Teixeira, L.S.; Dinali, L.A.F.; Simões, N.S.; de Souza Borges, W.; Borges, K.B. Novel restricted access material combined to molecularly imprinted polymer for selective magnetic solid-phase extraction of estrogens from human urine. Microchem. J. 2019, 149, 104043. [Google Scholar] [CrossRef]
  110. de Oliveira, H.L.; Teixeira, L.S.; Dinali, L.A.F.; Pires, B.C.; Simões, N.S.; Borges, K.B. Microextraction by packed sorbent using a new restricted molecularly imprinted polymer for the determination of estrogens from human urine samples. Microchem. J. 2019, 150, 104162. [Google Scholar] [CrossRef]
  111. Bousoumah, R.; Antignac, J.P.; Camel, V.; Grimaldi, M.; Balaguer, P.; Courant, F.; Bichon, E.; Morvan, M.-L.; Le Bizec, B. Development of a molecular recognition based approach for multi-residue extraction of estrogenic endocrine disruptors from biological fluids coupled to liquid chromatography-tandem mass spectrometry measurement. Anal. Bioanal. Chem. 2015, 407, 8713–8723. [Google Scholar] [CrossRef]
  112. Li, Z.; Wang, J.; Chen, X.; Hu, S.; Gong, T.; Xian, Q. A novel molecularly imprinted polymer-solid phase extraction method coupled with high performance liquid chromatography tandem mass spectrometry for the determination of nitrosamines in water and beverage samples. Food Chem. 2019, 292, 267–274. [Google Scholar] [CrossRef]
  113. Arabi, M.; Ostovan, A.; Bagheri, A.R.; Guo, X.; Wang, L.; Li, J.; Wang, X.; Li, B.; Chen, L. Strategies of molecular imprinting-based solid-phase extraction prior to chromatographic analysis. TrAC Trends Anal. Chem. 2020, 115923. [Google Scholar] [CrossRef]
  114. Zhu, G.; Fan, J.; Gao, Y.; Gao, X.; Wang, J. Synthesis of surface molecularly imprinted polymer and the selective solid phase extraction of imidazole from its structural analogs. Talanta 2011, 84, 1124–1132. [Google Scholar] [CrossRef] [PubMed]
  115. Madikizela, L.M.; Tavengwa, N.T.; Chimuka, L. Applications of molecularly imprinted polymers for solid-phase extraction of non-steroidal anti-inflammatory drugs and analgesics from environmental waters and biological samples. J. Pharm. Biomed. Anal. 2018, 147, 624–633. [Google Scholar] [CrossRef] [PubMed]
  116. Madikizela, L.M.; Ncube, S.; Chimuka, L. Green chemistry features in molecularly imprinted polymers preparation process. In Comprehensive Analytical Chemistry; Elsevier: Amsterdam, The Netherlands, 2019; Volume 86, pp. 337–364. ISBN 0166-526X. [Google Scholar]
  117. Yang, B.; Fu, C.; Li, J.; Xu, G. Frontiers in highly sensitive molecularly imprinted electrochemical sensors: Challenges and strategies. TrAC-Trends Anal. Chem. 2018, 105, 52–67. [Google Scholar] [CrossRef]
  118. Garcia, R.; Gomes da Silva, M.D.R.; Cabrita, M.J. “On-off” switchable tool for food sample preparation: Merging molecularly imprinting technology with stimuli-responsive blocks. Current status, challenges and highlighted applications. Talanta 2018, 176, 479–484. [Google Scholar] [CrossRef]
  119. Vaneckova, T.; Bezdekova, J.; Han, G.; Adam, V.; Vaculovicova, M. Application of molecularly imprinted polymers as artificial receptors for imaging. Acta Biomater. 2020, 101, 444–458. [Google Scholar] [CrossRef]
  120. Gui, R.; Jin, H.; Guo, H.; Wang, Z. Recent advances and future prospects in molecularly imprinted polymers-based electrochemical biosensors. Biosens. Bioelectron. 2018, 100, 56–70. [Google Scholar] [CrossRef]
Chemosensors 09 00151 g001 550
Figure 1. Chemical structures of hormone group representatives.
Figure 1. Chemical structures of hormone group representatives.
Chemosensors 09 00151 g001
Chemosensors 09 00151 g002 550
Figure 2. General scheme for molecular imprinting (figure adapted from [64]).
Figure 2. General scheme for molecular imprinting (figure adapted from [64]).
Chemosensors 09 00151 g002
Chemosensors 09 00151 g003 550
Figure 3. General synthetic scheme for polymerization.
Figure 3. General synthetic scheme for polymerization.
Chemosensors 09 00151 g003
Chemosensors 09 00151 g004 550
Figure 4. General preparation scheme for surface imprinting on a nanoparticle (adopted from Riedel and Mizaikoff 2019 [79] with permission).
Figure 4. General preparation scheme for surface imprinting on a nanoparticle (adopted from Riedel and Mizaikoff 2019 [79] with permission).
Chemosensors 09 00151 g004
Chemosensors 09 00151 g005 550
Figure 5. Magnetic MIP–SPE of steroid hormones reported by Guc and Schroeder [64].
Figure 5. Magnetic MIP–SPE of steroid hormones reported by Guc and Schroeder [64].
Chemosensors 09 00151 g005
Chemosensors 09 00151 g006 550
Figure 6. Zipper-like on/off switchable magnetic molecularly imprinted microspheres used by Zhu and colleagues [104] for the solid-phase extraction of steroid hormones in milk samples. (A) Represent the preparation of molecularly imprinted microspheres and (B) on/off-switchable recognition mechanisms of molecularly imprinted microspheres.
Figure 6. Zipper-like on/off switchable magnetic molecularly imprinted microspheres used by Zhu and colleagues [104] for the solid-phase extraction of steroid hormones in milk samples. (A) Represent the preparation of molecularly imprinted microspheres and (B) on/off-switchable recognition mechanisms of molecularly imprinted microspheres.
Chemosensors 09 00151 g006
Chemosensors 09 00151 g007 550
Figure 7. Surface-imprinted magnetic polymers used by Du and coworkers [107] for the extraction and determination of dexamethasone.
Figure 7. Surface-imprinted magnetic polymers used by Du and coworkers [107] for the extraction and determination of dexamethasone.
Chemosensors 09 00151 g007
Table
Table 1. Global hormone concentration levels.
Table 1. Global hormone concentration levels.
CountrySample TypeType of HormoneConc. LevelRef.
MalaysiaWastewater17-β-E2 and 17-α-EE20.02–93.9 ng/L[29]
South AfricaWastewater17-β-E215–2000 ng/L[9]
BrazilRiver water17-β-E214.9 µg/L[30]
ChinaRiver and wastewaterE1 and E32.1–360 ng/L[31]
FranceButterMedroxyprogesterone4.1 µg/kg[32]
PortugalWastewater17-β-E20.085 µg/L[33]
RomaniaWastewaterE3 and ethynylestradiol (EE)2.6–4.7 µg/L[34]
PolandGround waterE1309 ng/L[35]
ChinaSoilPRO, androstenedione, TST, and 17α-E20.06–1/20 µg/kg[36]
HungaryRiver waterE2, coprostanol, cholesterol, stigmasterol, and β-stosterol0.322–488 µg/L[37]
ChinaRiver waterPRO, boldenone, and norgestrel8.22–66.2 ng/L[38]
ChinaRiver and surface waterE1, E3, and bisphenol A (BPA)1.0–690 ng/L[39]
SwitzerlandMilkE1, PRO, hydroprogesterone, cortisone, 4-androstenedione, and E210–342 ng/kg[40]
BrazilSurface waterE1, 17β-E2l, PRO, and 17α-EE20–5.84 µg/L[41]
South Africa Wastewater and river waterPRO, E1, E2, and E30–7133 ng/L[42]
SerbiaSurface and wastewaterCholesterol, coprostanol, campesterol, stigmasterol, β-sitosterol, and sitostanol12–4808 ng/L[43]
Chile Human urineTST, PRO, and E20.20–21.23 ng/L[44]
Table
Table 2. Summary of application of MIPs for the determination of hormones in water samples.
Table 2. Summary of application of MIPs for the determination of hormones in water samples.
HormonesMatrixAnalytical TechniquesPolymerization MethodLOD (µg/L)%RecoveryReference
17β-E2Lake, river water, effluentHPLCOne-spot solvothermal reaction0.0488.3–99[47]
E1, E3, 17β-E2, 17α-EE2, trans-androsterone, TST, and PROWastewaterGC × GC‒TOFMS and LC‒MSBulk polymerization--[48]
Diethylstilbestrol (DES), E1, and E3River, lake and tap waterHPLC–UVSemicovalent polymerization 10–1696–98[49]
E3 and 17β-E2WastewaterLC–Q-TOFMS---[50]
E2, E3, and EE2Tap, drinking, river waterHPLC–FLDSurface polymerization2.5–5.872–102[51]
E2, E3, and DESLake and river waterHPLC–UVSurface polymerization and sol–gel method0.08–0.2785–95[52]
E1, E2, E3, DES, and EE2River waterLC–MS-0.0045–0.009847–104[53]
E1, E2, and DESLake and river waterHPLC–DAD-0.3–1.575–93[54]
E1, E2, E3, and EE2WastewaterHPLC-1.96–2.7681–113[55]
17α-E2, 17β-E2, E1, hexestrol (HEX), 17α-EE2, DES, dienestrol (DS), zearalenone (ZEN), α-zearalanol (α-ZAL), and β-zearalanol (β-ZAL)Mineral water and wastewaterHPLC–DAD-0.01–0.4465–101[56]
17β-E2, E1, and E3WaterHPLC–DAD/ECDBulk polymerization 0.07–10.9974–82[57]
E2, EE, DES, ethisterone (ES), and E1River waterHPLC–UVSurface polymerization0.1 to 0.26 mmol/L50–96[58]
estrogen dienestrol (DIS)SeawaterHPLC–DADSurface polymerization0.1687.3–96.4[59]
GC × GC–TOFMS = comprehensive two-dimensional gas chromatography–time-of-flight mass spectrometry, LC–MS = liquid chromatography–mass spectroscopy, HPLC–PDA = high-performance liquid chromatography–photodiode array, UV = ultraviolet spectrophotometry, LC–Q-TOFMS = liquid chromatography–quad time-of-flight mass spectroscopy, HPLC–DAD = high-performance liquid chromatography–diode array detector, ECD = electrochemical detector, HPLC–FLD = high-performance chromatography–fluorescence detection.
Table
Table 3. Summary of application of MIPs for the determination of hormones in food samples.
Table 3. Summary of application of MIPs for the determination of hormones in food samples.
HormonesFood TypeAnalytical TechniquePolymerization MethodLOD µg/L%RecoveryRefs
E2Goat milkHPLC–PDABulk polymerization4.8176–90[94]
PRO, TST, β-sitosterol, cholesterol, and campesterolWhite meat, egg yolks, and vegetablesHPLC-0.003–0.00597–101[95]
E2Milk powderUVSurface polymerization9.53384[96]
E2Milk HPLC–UVSurface polymerization0.0189–92[97]
E2BeefHPLC–PDASurface initiated atom transfer radical polymerization0.2597–99[98]
E1, E2, and E3Milk HPLCSurface polymerization-81.6–91.6[99]
E1, E3, and EE2Fish and shrimpHPLC–UVMultiple copolymerization0.98–2.3980–94[92]
E1, E2, E3. and DESMilk powder HPLC–UVsurface polymerization1.5–5.5 ng/g81–95[100]
E2 and E3Milk tabletsHPLC–UVSurface polymerization1.49–1.8389.1–93.5[101]
DESPork and chickenHPLC–UVSurface polymerization0.28–0.4783–99[102]
E2 MilkHPLC–FLD-0.00695–107[103]
HPLC–PDA = high-performance liquid chromatography–photodiode array, UV = ultraviolet spectrophotometry, HPLC–FLD = high-performance chromatography–fluorescence detection.
Table
Table 4. Summary of application of MIPs for the determination of hormones in biological samples.
Table 4. Summary of application of MIPs for the determination of hormones in biological samples.
HormonesMatrixAnalytical Techniques Polymerization MethodLOD µg/L%RecoveryRefs
E1 and E3UrineHPLC–UV-25–3270–80[109]
EE2 and E2Urine HPLC–UV-76–9296–99[110]
E1, 17α- α-E2, β-E2, E3, EE2, DES, BPA, bisphenol S (BPS), 4-n-octylphenol (OP), 4-n-coumestrol (COU), genistein (GEN), and enterolactone (ENT)Maternal serum, cord serum, and urineUHPLC–MS/MS-0.01–0.7>100[111]
PRO and TSTHuman urineHPLC–DADBulk polymerization0.47>80[111]
HPLC–UV = high-performance liquid chromatography with ultraviolet, UHPLC–MS/MS = ultrahigh-performance liquid chromatography coupled with tandem mass spectroscopy, DAD = diode array detector.
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Mpupa, A.; Selahle, S.K.; Mizaikoff, B.; Nomngongo, P.N. Recent Advances in Solid-Phase Extraction (SPE) Based on Molecularly Imprinted Polymers (MIPs) for Analysis of Hormones. Chemosensors 2021, 9, 151. https://doi.org/10.3390/chemosensors9070151

AMA Style

Mpupa A, Selahle SK, Mizaikoff B, Nomngongo PN. Recent Advances in Solid-Phase Extraction (SPE) Based on Molecularly Imprinted Polymers (MIPs) for Analysis of Hormones. Chemosensors. 2021; 9(7):151. https://doi.org/10.3390/chemosensors9070151

Chicago/Turabian Style

Mpupa, Anele, Shirley Kholofelo Selahle, Boris Mizaikoff, and Philiswa Nosizo Nomngongo. 2021. "Recent Advances in Solid-Phase Extraction (SPE) Based on Molecularly Imprinted Polymers (MIPs) for Analysis of Hormones" Chemosensors 9, no. 7: 151. https://doi.org/10.3390/chemosensors9070151

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop