Precipitation polymerization: a versatile tool for preparing molecularly imprinted polymer beads for chromatography applications

Abstract

Precipitation polymerization is a robust method for synthesis of polymers in the form of micro- and nanospherical beads. This spherical particulate polymer morphology is advantageous for several applications of molecularly imprinted polymers (MIPs) including their utilization as sorbents in chromatography and solid phase extraction (SPE), in sensing and drug delivery applications. In this context, this review aims to compile and present recent advances and outcomes of chromatography and SPE applications of MIPs prepared by precipitation polymerization for recognition of target analytes in complex matrices such as environmental, food and biological samples. First, the basic concept of MIPs synthesis by using precipitation polymerization is introduced. This is followed by details on the chromatography and SPE applications of MIPs prepared via precipitation polymerization. The controlled radical precipitation polymerization and other advances in the field are introduced. Finally, the review is concluded with the prospects for future research.

---

1. Introduction

Molecularly imprinted polymers (MIPs) are synthetic materials with template-induced binding sites to recognize the analyte of interest (template molecule) in preference to other structural analogues.^1,2 MIPs are synthesized by copolymerization of the functional monomer and the cross-linker in the presence of the template molecules. Removal of the template molecules from the highly cross-linked polymer matrixes results in the formation of recognition cavities that are complementary to the template molecules in shape, size and chemical functionality. These recognition cavities can selectively rebind the template molecules from the complex matrix.³

MIPs are one of the most intensively studied materials during past two decades owing to their obvious advantages of predictable specific recognition, comparatively low cost, easy preparation, chemical stability to harsh chemical and physical conditions and excellent reusability.^2–4 MIPs have potential applications in various fields such as affinity separation,⁵ chemical sensors,⁶ drug delivery⁷ and enzyme-like catalysis.⁸ Chromatography and solid-phase extraction (SPE) are the major areas of application of MIPs, wherein it is used as sorbent in chromatography columns and SPE cartridges due to its selectivity for target analyte.^9,10

MIPs are synthesized in a variety of physical forms such as monoliths, porous microspheres, thin films, nanostructured materials and hydrogels¹¹ or immobilized over a solid silica support according to the demand of the intended application.¹² Most of the MIPs are prepared by bulk polymerization where, the monomer and the crosslinker are dissolved in the minimum quantity of porogenic solvent.^3,10 Indeed many MIPs have been prepared with unprecedented selectivity by utilizing this generic concept which significantly facilitated their use for the intended application.^8,13 In bulk polymerization, the resultant polymer monolith requires a tedious process of crushing, grinding and sieving to obtain the desired sized particles, moreover this also leads to wastage of useful fraction of MIPs.^3,11,14,15

It is noteworthy to mention that spherical and monodispersed microspherical particles are more desirable for numerous applications, particularly when the MIPs are indented to use as stationary phases (sorbents) in SPE or chromatography column.¹⁰ This is significant in order to circumvent the problems such as slow mass transfer of analyte, generation of backpressure in chromatography columns and peak tailing in resulting chromatograms which occur due to the irregularly shaped MIPs particles synthesized by bulk polymerization.^{3,9,10,16–19}

Many successful methods are developed to synthesize uniformly sized, spherical MIPs beads such as suspension polymerization, emulsion polymerization, solution polymerization and multi-step swelling polymerization.^11,19,20 Some merits and limitations are associated with these methods as presented in Table 1.^11,15

Table 1 Merits and limitations of polymerization methods used to synthesize the MIPs^11,15

Polymerization method	Merits	Limitations
Bulk polymerization	Simplicity of method makes it possible to perform in any laboratory	Tedious procedures of grinding and sieving
	Do not require particular skills or sophisticated instrumentation	Wastage of useful polymer fraction, irregularly sized particles
Suspension polymerization (either in aqueous phase or in perfluorocarbon continuous phase)	Spherical particles, highly reproducible results, large scale synthesis is possible	Phase partitioning complicates the system
	Use of regular surfactant is allowed in aqueous continues phase	Water is incompatible solvent for some of the imprinting procedures
	Applicable to most imprinting systems, particle size can be adjusted	Not applicable to imprinting using hydrophilic monomers
		Specialized surfactant required for perfluorocarbon continuous phase
Multi-step swelling polymerization	Monodispersed beads of controlled diameter	Complicated procedures and reaction conditions
	Excellent particle size for chromatography applications	Need for aqueous emulsions
		Not compatible with non-covalent imprinting system
Emulsion polymerization	Allows to predetermine the polymer particle size	Complicated processes and use of stabilizers and surfactants can contaminate the MIPs
	Formation of spherical particles	Stabilizers and surfactants are difficult to remove from the resulting MIPs and may generally interfere with the imprinting procedure

---

Precipitation polymerization is a method of preparation of polymeric microspheres with sufficient control of product morphology. It is a facile polymerization method which gives rise to microspheres with clean and smooth surfaces and suitable particle sizes.^3,17–20 The advantages of this method include simplicity of preparation, lack of need of stabilisers or other additives, compatibility with high degrees of crosslinking agents and use of polar aprotic solvents as porogen which are capable of preserving non-covalent interactions between template and monomer. These advantages have made this method ‘the widely employed preparation method’ for the MIPs microspheres in molecular imprinting.²¹ Precipitation polymerization is also applied in numerous other fields such as for the preparation of polymer particles with interesting internal structure, electrophoretic displays and environmentally responsive polymers.²²

Precipitation polymerization method was developed by Stover et al.²³ and application of this method to MIPs was first proposed by Ye et al.¹⁵ for the synthesis of MIPs beads in the submicrometer range.

Several excellent reviews are published covering various aspects of molecular imprinting and MIPs.^{2,3,7,9,24–34} This minireview is especially devoted to the chromatography and SPE applications of MIPs prepared using precipitation polymerization. In this work, research articles reporting chromatography and SPE application of MIPs prepared via precipitation polymerization during past five years are included. It is a hope that this minireview will serve helpful for future research on the precipitation polymerization based MIPs.

2. Precipitation polymerization – a robust way to prepare MIPs beads

A pre-requisite towards better understanding of the subject makes it essential to discuss the chemistry of precipitation polymerization, its development and present status of the field of molecular imprinting. Precipitation polymerization is a surfactant-free heterogeneous polymerization process which begins initially as a homogeneous solution of crosslinker and monomer (typically <5% w/v) in a mixture of solvent and porogen (step A Fig. 1).^5,18,20,21 As the polymerization starts, oligomers and nuclei are being formed, the oligomers are still soluble in the medium (solvent), the nuclei precipitate resulting in a heterogeneous mixture. The nuclei are swollen by porogen and the continuous phase becomes less rich in porogen. The nuclei are stabilized by a layer of oligomers (step B in Fig. 1).³⁵

Fig. 1 Schematic description of the stages of precipitation polymerization for formation of porous MIPs beads. (A) Initially, monomer, crosslinker and initiator are dissolved in the mixture of solvent and porogen resulting in formation of a homogeneous system. (B) Growth of oligomers and nuclei takes place by radical polymerization. (C) Nuclei grow by adding fresh molecules of monomers and oligomers from the medium resulting in formation of polymer segregates (porogen is shown by yellow colour in the schematic).

As the polymerization continues the nuclei do not overlap but only grow by adding fresh monomers and oligomers from the continuous phase.³⁵ The porogen that is initially absorbed by the growing nuclei, separates from the particle and forms the pores (step C in Fig. 1). In summary, during polymerization the growing polymer segregates from the solution while continuously capturing and consuming monomers and oligomers, eventually forming beads in the micro or sub-micrometer range under proper conditions.^16,20 Near monodisperse, spherical particles can be routinely prepared in good yields via this method.^17,19,21

2.1 Selection of precursors for synthesis of MIPs using precipitation polymerization

Selection of appropriate precursors is a crucial step in the synthesis of MIPs. The functional monomers are responsible for providing functionalities which form a complex with the template either by covalent or non-covalent interactions. The strength of the interactions between the template and monomer determines the affinity and selectivity of recognition sites in MIPs. A stronger interaction results in stable template–monomer complex which in turn provides higher binding capacity in the MIPs. Generally, selection of the functional monomers is based on its ability to form non-covalent interactions with the template molecules viz. ionic bonds, hydrogen bonding interaction. Commonly used functional monomers include methacrylic acid (MAA), acrylic acid (AA), 2- or 4-vinylpyridine (2-VP or 4-VP), acrylamide (AM), trifluoromethacrylic acid (TFMAA) and hydroxyethyl methacrylate (HEMA). Spectroscopic measurements and computer simulation are also used for selection of an optimized functional monomer.³⁶

The crosslinker is an important precursor of MIPs which exhibits at least two polymerizable double bonds in its structure. The role of the crosslinker is to fix template–monomer complex and thereby form highly cross-linked rigid polymer network. Types and amounts of cross-linkers have profound influence on selectivity and binding capacity of the MIPs. Commonly used cross-linkers involve ethylene glycol dimethacrylate (EGDMA), trimethylolpropane trimethacrylate (TRIM), N,N-methylenebisacrylamide (MBAA) and divinylbenzene (DVB).

In precipitation polymerization, all the precursors involved in the synthesis of the MIPs are highly diluted in a solvent (or a mixture of two solvents). This solvent is able to solubilise all the precursors at the temperature of polymerization; it is also able to solubilise the oligomers formed up to a certain extent. This temperature is known as theta temperature and the solvents with this particularity for a given polymer are known as theta solvents. Once the polymer reaches a certain critical mass, the solvent is no longer able to hold the polymeric chain in the solution thus precipitating particles off the solution. Therefore, selection of a proper theta solvent is essential to obtain discrete particles.

Acetonitrile, being a theta solvent has predominantly used continuous phase in precipitation polymerization.²⁴ It is responsible for precipitating the polymeric chains once they reach a certain critical mass and it also avoids the gel-formation. Frequently, a mixture of acetonitrile–toluene (75 : 25 v/v) is used when the main goal is to create porous microspheres with high surface area. Porosity in MIPs is crucial as it enables easy ‘in and out’ diffusion of template molecules during the process of binding and extraction. Toluene is widely used for this purpose as the porogen. It brings porosity to the polymer and keeps the polymeric chains growing in the solution from collapsing.³⁷ Apart from the aforementioned systems; many solvents and solvent mixture have been used as a continuous medium in precipitation polymerization.^22,38

Nature of precursors influences morphology of the MIPs. According to the mechanism of precipitation polymerization, the growing nuclei are swollen by the solvent and capture soluble oligomers continuously from solution.²³ Thus, matching the solubility parameters of precursors, particularly solvent and polymer is important to obtain desired particle size and porosity in the resultant MIPs. Any mismatch between the solubility parameter of the solvent and the polymer leads to a early phase separation of polymer and results in the MIPs with negligible porosity and very low specific surface areas.³⁹ Whereas, a better match between the solubility parameter of the solvent and the polymer leads to better solvation of the polymer with well-developed pore structure and high specific surface areas. The Hildebrand solubility parameter (δ), corresponding to the cohesive energy density of solvent and polymers, is used to interpret the role of solvents in polymerization reactions. Other porogen systems, such as methanol–water, chloroform–methanol and THF can be used however; nature of functional monomer and crosslinker should be also considered.

The precipitation polymerization yield the particulates typically in microns or even in nanometer range which are useful particularly in capillary electro chromatography^5,14,38,40 and drug delivery applications.^41–44 However, the particle size in the range of 3–5 μm range is generally used for MIPs as stationary phase in chromatography. An excellent discussion of all the parameters affecting the polymerization process was reported by Wang et al.²⁰ in which they fully detailed the influence of each parameter involved in the polymerization process. Mosbach et al.¹⁷ reported that by polymerizing a mixture of DVB and TRIM in different ratios, it was possible to tune the size of the resulting MIPs particles in the range 130 nm to 2.4 μm. These pioneering investigations triggered an intensive use of precipitation polymerization in the recent years. Variety of solvents systems, monomers and experimental conditions are used to synthesize monodisperse MIPs microspheres with clean and smooth surfaces and to study the effect of synthetic conditions on the morphology of MIPs.^{5,19,23,38,45,46}

The spherical microbeads of MIPs produced using precipitation polymerization (hereafter mentioned as MIPs-PP) have been successfully applied to imprinting and recognition of large variety of target molecules for various applications.^47–52 The advantages of this method in the molecular imprinting include quick delivery of high quality imprinted polymers with minimal optimization in a one-step process in good yield.^5,17 The uniqueness of the precipitation polymerization is development of high level of crosslinking in individual particle. The coagulation between the particles is prevented due to the hard and resilient nature of each particle.³⁵ In comparison with the MIPs prepared by conventional bulk polymerization, the MIPs-PP generally proved superior in terms of specific affinity for the template.¹⁸

3. Chromatography and SPE applications of MIPs-PP

After making initial introduction and background of the subject in first two sections, this section put forwards a proper focus on chromatography and SPE application of MIPs-PP. The prevailing trend in chromatography is use of shorter and narrower columns packed with sorbent microspheres which can withstand extreme pressure.⁵³ Since, MIPs-PP are highly cross linked, rigid and chemically resistant to high temperature and pressure, this makes them suitable support in chromatographic columns and chemically more stable alternative to the commonly used silica and bonded silica materials such as C8 or C18. Molecularly imprinted solid phase extraction (MISPE) is applied for the cleanup, enrichment and preconcentration of variety of target compounds.^9,11 This section presents MISPE applications of MIPs-PP and it is divided in to the further subsections according the nature of the target analyte and the matrix from which it is extracted.

3.1 MIPs-PP for recognition and extraction of pharmaceutically active molecules from complex herbal matrices

Natural products such as alkaloids, flavones and polyhydric phenols exhibit several medicinal properties viz. antioxidants, anticancer, antibiotic, anti-inflammatory and are known to provide protection from the damaging effects of the free radicals. These compounds are of enormous interest to pharmaceutical industries due to their pharmaceutically active properties. Since, these compounds are present in ample quantity in the natural herbal matrices; their extraction is preferred. The traditional extraction methods are non-selective in nature moreover; the complexity of herbal matrices further complicates the process. The MIPs attracted considerable attention as SPE sorbents for selective extraction of pharmaceutically active compounds from complex herbal extracts.⁵⁴ Many alkaloids, terpenoids and polyphenolics have been extracted from the complex plant samples using the MIPs-PP (Fig. 2).

Fig. 2 Chemical structures of natural products imprinted via precipitation polymerization.

The MIPs-PP were prepared for the terpenoids (andrographolide and dehdroandrographolide) for their enrichment and preconcentration from the herb Andrographis paniculata.⁵⁵ The capacity of MIPs to bind the target analytes was found to be twice in comparison with conventional C18 material in SPE. Similarly, MIPs-PP was prepared for the extraction of protocatechuic acid from the Chinese herbal medicine.⁵⁶ The MIPs-PP microsphere exhibited good selectivity and extraction performance that resulted in about 82% recovery of target protocatechuic acid from the herbal extract in an optimized MISPE protocol. It is important to mention that these reports were focused on the development of an optimized MISPE protocols for preconcentration and isolation of target compounds nevertheless less attention was given to the study of MIPs-PP morphology.

There are some reports, in which the link between MIPs-PP particle morphology with its molecular recognition abilities was established. Caffeic acid MIPs-PP resulted in formation of monodispersed spherical microspheres of 5 to 1.5 μm particle diameters and high specific surface area of 340 m² g⁻¹.⁵⁷ The MIPs-PP microspheres were found to be selective for target caffeic acid in presence of its other eight structural analogues and achieved nearly 81% recovery in spiked apple juice samples without performing any sample clean-up steps when used as stationary phase in HPLC column. MIPs-PP for gallic acid, prepared in the 100% acetonitrile resulted in the formation of microspheres (Fig. 3a) with particle size approximately 4 μm and low surface area of 96.73 m² g⁻¹. Whereas, MIPs-PP prepared in a mixture of acetonitrile–toluene (75 : 25 v/v) (Fig. 3b) resulted in formation of nanoparticles (0.8–1000 nm) and high surface area of 345.9 m² g⁻¹.⁵⁸ The results showed that effect of toluene on the particle size of MIPs depends on the solubility parameter of crosslinker EGDMA. MIPs-PP prepared in the 100% acetonitrile extracted about 75% of pure gallic acid whereas; MIPs-PP prepared in a mixture of acetonitrile–toluene (75 : 25 v/v) extracted about 83% pure gallic acid from the complex matrix of Emblica officinalis fruits. The MIPs-PP for the templates matrine and oxymatrine with particle size of about 3 μm were found to be highly selective and resulted in much cleaner HPLC chromatograms of concentrated extract of herbs compared to the conventional C18 cartridges.⁵⁹ The template chlorogenic acids being hydrophilic in nature did not dissolve in conventional porogens, therefore, it was first dissolved in small portion of polar co-solvents such as MeOH and DMSO for preparation of MIPs-PP.⁶⁰

Fig. 3 SEM of (a) MIP prepared in 100% acetonitrile and (b) MIP prepared in mixture of acetonitrile–toluene (75 : 25 v/v).

The particle size of the resulting polymers were about 4 μm and it could extract the target from the herbal sample.

Synthesis of MIPs-PP generally involve a longer polymerization time of 24 to 48 hours with the conventional thermal methods. MIPs-PP was prepared for the templates kaempferol⁶¹ and podophyllotoxin⁶² using microwave induced precipitation polymerization. This method required much shorter polymerization time of 150 minutes compared to the conventional thermal method and resulted in better particle morphology. Moreover, MIPs-PP also exhibited high selectivity and achieved nearly 90% extraction of targets from traditional Chinese medicines.

Use of ionic liquid as monomer is emerging in the field of polymer science. Various ionic liquid (1-vinyl-3-carboxymethylimidazolium bromide, 1-vinyl-3-carboxyethylimidazolium bromide, 1-vinyl-3-carboxybutylimidazolium bromide, or 1-vinyl-3-carboxypentylimidazolium bromide) were used as functional monomers to imprint an alkaloid synephrine and the resultant MIPs-PP exhibited good selectivity and binding capacities.⁶³ These results indicated that ionic liquid as a functional monomer can provide a wide choice of monomers for synthesis of MIPs.

In another approach, the molecular imprinting was clubbed with supercritical fluid extraction technique to improve the specificity of crystallization of products.⁶⁴ Supercritical fluids are of great interest for the crystallization of compounds which are difficult to separate. Oleanolic acid MIPs-PP was placed in the form of thin-layer plates in a supercritical fluid extractor. The MIPs-PP induced crystallization of oleanolic acid reached nearly 93%, with the purity of 95%. The details of precursors and polymerization method used for synthesis of aforementioned MIPs-PP are presented in Table 2. A successful application of MIPs-PP for extraction of natural compounds is established in the literature. It is noteworthy to mention that the template molecule has a remarkable influence on the morphology of resultant polymer product.⁶⁵ Template molecules of natural origin exhibits many polar groups in their structure (as shown in Fig. 2) thus, it is necessary to study the influence of these template molecules on the polymer morphology and the yield of resulting MIPs.

Table 2 Details of the precursors and polymerization method used for synthesis of MIPs-PP for pharmaceutically active molecules

Template	Functional monomer and crosslinker	Porogen	Method of polymerization	Ref.
Andrographolide and dehdroandrographolide	Acrylamide–EGDMA	Acetonitrile : toluene (75 : 25 v/v)	Thermal	55
Protocatechuic acid	Acrylamide–EGDMA	Acetonitrile	Thermal	56
Caffeic acid	4-Vinylpyridine–DVB	Acetonitrile : toluene (75 : 25 v/v)	Thermal	57
Gallic acid	Acrylic acid–EGDMA	Acetonitrile : toluene (75 : 25 v/v)	Thermal	58
Matrine and oxymatrine	Methacrylic acid–DVB	Acetonitrile : toluene (75 : 25 v/v)	Thermal	59
Chlorogenic acid	Methacrylic acid–DVB	Acetonitrile : toluene (75 : 25 v/v)	Thermal	60
Kaempferol	2-Vinylpyridine–EGDMA	Acetonitrile : methanol (80 : 20 v/v)	Microwave	61
Podophyllotoxin	Acrylamide–EGDMA + DVB	Acetonitrile	Microwave	62
Synephrine	Ionic liquids + EGDMA	Methanol : water (4 : 1 v/v)	Thermal	63
Oleanolic acid	Acrylamide	Chloroform : methanol (3 : 1 v/v)	Thermal	64

---

3.2 MISPE of drugs from biological samples using MIPs-PP

The detection of drugs and regulated substances is of prime interest in biomedical fields and forensics. This includes monitoring drug safety level in medicinal compositions to address to toxicological issues and detection of illicit consumption of drugs to prove involvement of the regulated substances in crimes and drug doping in sports.⁶⁶ The drugs and their metabolites are routinely monitored in human body fluids such as serum and urine for the aforementioned purposes. Low concentration of drugs and their metabolites and complex nature of the matrix creates difficulty in selective detection procedures therefore the MIPs are implemented in these areas of applications due to the prime advantage of selectivity.

MIPs-PP were synthesized using MAA and DVB for Emtricitabine, an anti-HIV drug.⁶⁷ The effect of chloroform as dispersant on the morphology of MIPs was observed. Uniformly sized MIPs-PP of 3 to 5 μm was formed with the 1 : 3 (v/v) ratio of chloroform to acetonitrile. The MIPs-PP exhibited good selectivity and above 90% recovery of emtricitabine was achieved in human serum samples. Aconitine MIPs-PP was prepared using MAA and EGDMA in varying composition of different porogens.⁶⁸ The SEM revealed spherical MIPs particles with diameters ranging from 1–2 μm in the porogen toluene. The MIPs-PP exhibited high selectivity for aconitine in presence of other poisonous alkaloids and could extract nearly 89% aconitine from spiked serum samples. Tripterine MIPs-PP with the average particle size around 800 nm was prepared using MAA : EGDMA (50 : 50 v/v) in porogen ethyl acetate.⁶⁹ The MISPE revealed good selectivity and exhibited a recovery up to 95% of tripterine from spiked urine sample. Different ratio of acetonitrile and toluene was used in the synthesis of MIPs-PP for bisphenol-A in order to study their influence on the morphology of the MIPs-PP microspheres.⁷⁰ MISPE of urine sample resulted in high selectivity for bisphenol A in presence of its structural analogues with recoveries up to 95%.

There are some reports where MIPs-PP exhibited some cross-reactivity for other structural analogues present in the matrix. However, this cross-reactivity was utilized as an advantage for simultaneous extraction of multi targets using group selective MIPs. Spherical and monodispersed barbital MIPs-PP of about 4 μm size was synthesised using a designer functional monomer 2,6-bis-acrylamidopyridine and DVB-80 as the crosslinker in acetonitrile : toluene 4% (w/v).⁷¹ These MIPs-PP showed high binding capacity for the template barbital as well as some useful cross-selectivity for other barbiturates. The percent extraction of four barbiturates from human urine was 40% barbital, 65% phenobarbital, 47% secobarbital and 35% pentobarbital.

Computational modelling is used to select the functional monomer and porogen with high interaction energy for the templates in order to achieve high binding capacity and selectivity in MIPs.⁷² Some selective MIPs were developed using the functional monomer with the highest binding energy for the template in a computationally optimized procedure for the target drugs. Methocarbamol based MIPs-PP were computationally designed using semi-empirical and density functional theory (DFT) based quantum chemical calculation.⁷³ The AA and THF were found to be the best choices of functional monomer and porogen, respectively. Higher recovery of the drug methocarbamol was achieved in MISPE compared to the conventional C18 cartridge moreover, the cartridges filled with MIPs-PP were found to be reusable for clean-up of about fifteen plasma samples with no noticeable deterioration in performance. While conventional C18 cartridges lost their performance after one to two extractions. DFT based computational modelling was also applied for selection of best precursors for preparation of salbutamol MIPs-PP.⁷⁴

3.3 MIPs-PP for clean up, preconcentration and isolation of pharmaceuticals and toxicants from environmental and food samples

Development of detection and isolation methods for toxicants from the food and environmental matrices are gaining paramount importance as there is a widespread occurrence of chemical compounds, metals, drugs and their metabolites in the food and environmental samples in trace quantity. Persistent of pharmaceuticals and toxicants in the food and environment leads to several hazards ailments and toxicity issues in humans which may even lead to death. The presence of pharmaceuticals in the aquatic environment at low concentration levels are of great concern because of the potential impact on human health and the ecosystem. MIPs are successfully applied for clean-up and pre-treatment of aqueous samples before application of any analytical method as well as for direct detection and quantification of analytes without any pre-treatment.^75,76

MIPs-PP was synthesized for diclofenac (an important non-steroidal anti-inflammatory agent) using 2-VP, EGDMA and toluene.⁷⁷ Recoveries of diclofenac from water samples were about 95% moreover, the MIPs-PP were reusable for about 30 adsorption and desorption cycles. In the another report on diclofenac MIPs-PP synthesized by 2-VP, EGDMA and toluene, the sorption of diclofenac was very rapid and reached to an equilibrium within 15 min. The selectivity and adsorption capacity of MIPs-PP was not influenced even in the presence of the interfering humic acid up to its concentration of 10 mgL⁻¹. The MIPs-PP exhibited reusability of twelve times without significant loss in their performance.⁷⁸

Combination of novel crosslinkers with routinely used crosslinkers were also used in order to enhance the morphology and binding capacity of MIPs-PP. In the reports by Rodríguez et al.,⁷⁹ MIPs-PP microspheres were developed with monomer and crosslinker MAA, HEMA, EGDMA and DVB for simultaneous detection of six fluoroquinolone antimicrobials in water samples. MIPs-PP around 50 to 100 nm particle diameter were prepared for levofloxacin, using MAA and 2-ethyl-2-(hydroxymethyl)propane-1,3-diol as a crosslinker.⁸⁰ The MIPs exhibited higher binding capacity in acetonitrile compared to water with retention of selectivity. 4-Cumylphenol, an endocrine disrupter was imprinted using MAA and DVB in acetonitrile.⁸¹ The morphology investigations revealed the formation of monodispersed microspheres with the average size around 2 μm, high surface area above 400 m² g⁻¹ and high thermal stability. The MISPE of spiked water and soil samples method could detect 4-cumylphenol up to ng L⁻¹.

Beltran et al.⁸² prepared two MIPs by precipitation and bulk polymerization using esters of p-hydroxybenzoic acid (parabens) as templates. The synthesis of MIPs-PP was based on semi-covalent imprinting strategy whereas MIPs prepared by bulk polymerization was based on non-covalent molecular imprinting strategy. Authors reported that precipitation polymerization is a more suitable polymerization method to produce MIPs of suitable size for MISPE. Better packing of the cartridge was possible with the MIPs-PP than the irregular material derived from bulk polymerization. However, due to the use of the semi-covalent approach, a single interaction for the template was developed in MIPs-PP due to which it could not reach the expected recoveries in MISPE procedure. Thus, the binding performance of MIPs prepared by bulk polymerization was found better.

Foodstuffs are often contaminated with pharmaceuticals and pesticides residues in trace quantity which are difficult to analyze without any sample treatment since, the matrix is rather complex. The sample preparation greatly influences the reliability and accuracy of the detection method. MIPs are preferred over the conventional sorbents due to their high binding capacity and selectivity for analyte.^6,9 Tetracycline MIPs-PP were prepared by using MAA and TRIM in mixture of methanol and acetonitrile (30 : 35, v/v) for their detection in foodstuffs.⁸³ The MIPs-PP exhibited good imprinting efficiency for different tetracycline with recoveries above 90% (oxytetracycline, tetracycline, chlortetracycline and doxycycline). MIPs-PP for 17β-estradiol were prepared using TFMAA as functional monomer and TRIM by UV radiation induced polymerization.⁸⁴ MIPs-PP showed better selectivity and enrichment property than C18 with the recovery up to 85.5%. The rapid, specific and sensitive MISPE method for the foodstuffs was developed using these MIPs-PP compared with the conventional sorbents.

MIP-PPs were prepared for using MAA and EGDMA in butanone and N-heptane porogen for determination of dimethomorph in ginseng samples.⁸⁵ Similarly, MIPs-PP were prepared for dimethoate pesticides using methyl methacrylate and EGDMA in acetonitrile.⁸⁶ The SEM revealed monodispersed spherical beads with an average diameter of about 3 μm. MISPE procedure resulted in recoveries about 88% in the cucumber samples. Fluorescent MIPs-PP was prepared by copolymerization of AM with a small quantity of allyl fluorescein in the presence of cyhalothrin, a synthetic parathyroid pesticide.⁸⁷ The MIPs-PP microspheres exhibited high fluorescence intensity and high selective recognition for determination of cyhalothrin in honey.

Preservatives and colorant show toxic effects if present in quantities above their permissible limits. Also, the presence of heavy metal ions in food and environmental matrix is a cause of concern as these ions have chronic toxicity. MIP-PP were prepared using MAA and EGDMA for detection of 1,4-hydroxybenzoic acid esters preservatives in soybean sauce samples using a MISPE-HPLC.⁸⁸ The ionic liquid 1-butyl-3-methylimidazolium hexafluorophosphate ([Bmim][PF6]) was used as functional monomer with DVB as the cross-linker in a mixture of toluene and acetonitrile for synthesis of MIPs-PP for simultaneous isolation and determination of Sudan dyes from foodstuff samples.⁸⁹ The MIPs showed good binding capacity for the four sudan dyes compared with the traditional SPE adsorbents.

Molecular dynamics simulations in association with UV-spectrometric study were used to study the nature of interaction between the template and monomers. The optimized precursors were used for synthesis of malachite green dye MIPs-PP.⁹⁰ The resulting MIPs-PP revealed high potential to recognize the dye in presence of its structural analogues in MISPE.

Lead ion imprinted MIPs-PP were prepared using 4-VP and EGDMA in acetonitrile in presence of 4-(2-pyridylazo)resorcinol, a lead-binding ligand.⁹¹ The low detection limit was achieved for lead ion under the optimized conditions in MISPE of fruit juices and water samples. The details of detection limits and method of detection are presented in the Table 3.

Table 3 Application of MIP-PPs for analyzing compounds in complex sample matrices

Analytes	Sample matrices	Application	Detection techniques	Detection limit	Ref.
Emtricitabine	Serum sample	MISPE	HPLC	0.005 mg L^−1	67
Aconitine	Serum sample	MISPE	HPLC	0.0167 mg L^−1	68
Tripterine	Urine sample	MISPE	UV spectrophotometer	Not reported (N.R.)	69
Barbital	Urine sample	MISPE	Liquid chromatography	N.R.	70
Methocarbamol	Human plasma samples	MISPE	Differential pulse voltammetry, HPLC/UV	2.45 mg L^−1	72
Diclofenac	Tap water, river water and wastewater samples	MISPE	HPLC/DAD	690 mg L^−1	77
Fluoroquinolone	Water samples	Online MISPE	Liquid chromatography	0.0011 mg L^−1	80
4-Cumylphenol	Water samples, soil sample	MISPE	HPLC/DAD	0.0027 mg L^−1	81
Parabens	Water samples	MISPE	Liquid chromatography	N.R.	82
Tetracycline	Foodstuffs	MISPE	HPLC/mass spectrometry	0.001–0.003 mg kg^−1	83
17β-Estradiol	Milk powder	MISPE	HPLC	N.R.	84
Dimethomorph	Ginseng	MISPE	Gas chromatography	0.002 mg kg^−1	85
Dimethoate	Cucumber samples	MISPE	HPLC/UV	2.3 mg L^−1	86
Cyhalothrin	Honey samples	Optical sensing	Fluorescence spectrophotometer	0.004 nM	87
Sudan dyes	Foodstuffs	MISPE	HPLC	0.004–0.014 mg kg^−1	89
Lead ion	Water samples, fruit samples	MISPE	Flame atomic absorption spectrometry	0.009 mg L^−1	91

---

4. Controlled radical precipitation polymerization

Preparation of uniform crosslinked polymer microspheres; involves the direct preparation of functional polymer microspheres by copolymerizing functional monomers with crosslinkers via precipitation polymerization.³⁹ However, this requires a time-consuming optimization of polymerization parameters to obtain uniform polymer microspheres. The other method is by grafting the functional polymer layers onto the polymer particles prepared by precipitation polymerization which allows the facile grafting of all the functional monomers onto the surfaces of the preformed polymer particles, thus leading to their flexible preparation and more efficient functionalization.⁹²

Different polymerization mechanisms are developed for grafting of functional monomers, which are based on the conventional free radical polymerization, the ionic polymerization and controlled living radical polymerizations (CRP). CRP offers many advantages over the conventional free radical and ionic polymerization such as milder and less restricted reaction conditions as well as their applicability for a much larger range of monomers.⁹³ The controlled living radical precipitation polymerization (CRPP) approaches were developed by the introduction of CRP mechanism into the precipitation polymerization system, this includes atom transfer radical precipitation polymerization (ATRPP),^94,95 iniferter-induced “living” radical precipitation polymerization (ILRPP),⁹⁶ and RAFT precipitation polymerization (RAFTPP).^97–99 These polymerization methods are proven to be facile and highly efficient approaches not only for preparation of monodisperse, highly crosslinked and living polymer microspheres, but also for synthesis of MIPs. The thermodynamically controlled process of CRPP allows a more constant rate for the polymer chain growth, leading to homogeneous polymer networks and MIPs with homogeneous network structures can lead to improved binding properties.⁹³ An excellent review covering the various aspects of CRP is reported.⁹³

Bisphenol A⁹⁴ was imprinted using ATRPP and conventional precipitation polymerization. ATRPP resulted in MIP microspheres with much larger diameters and significantly higher high-affinity site densities in comparison with the MIPs sub-microspheres prepared via precipitation polymerization. This suggested that the application of ATRPP in the molecular imprinting field has great potential to improve the structural homogeneity and to enhance the template binding properties. The MIP microspheres prepared via ATRPP exhibited high binding capacity, fast template rebinding kinetics and an appreciable selectivity.

ILRPP proved to be a highly efficient approach for providing MIP microspheres with obvious molecular imprinting effects towards the template, fast template rebinding kinetics, and appreciable selectivity in presence of structurally related compounds. More importantly, ILRPP provided grafted polymer microspheres with enhanced dispersion stability in water at ambient temperature.⁹⁶

MIPs was prepared for 2,4-dichlorophenoxyacetic acid (2,4-D)-using RAFTPP and conventional precipitation polymerization.¹⁰⁰ RAFTPP provided MIP microspheres while only irregular MIP aggregates were obtained via conventional precipitation polymerization. The MIP microspheres prepared via RAFTPP exhibited improved binding capacity, larger binding constant with high-affinity sites and significantly higher binding site density in comparison with those prepared via conventional precipitation polymerization. This effect was ascribed to the controlled polymerization mechanism of RAFTPP which resulted in increased structural homogeneity and improved stability and integrity of the binding sites.

Chen et al.¹⁰¹ used RAFTPP to prepare atrazine imprinted MIPs. In comparison to the MIPs-PP, MIPs-RAFTPP exhibited uniform spherical particles with rough surfaces and significant amounts of micropores and high binding capacity. MIPs-RAFTPP were successfully utilized as SPE sorbent for the preconcentration and selective separation of atrazine in spiked lettuce and corn samples.

The RAFTPP is demonstrated to be a promising method for preparation of water compatible MIPs.^97–99 The RAFTPP is utilized for preparation of water-compatible MIP microspheres by the controlled grafting of ultrathin hydrophilic polymer shells onto the living MIP microspheres prepared by using surface-initiated RAFT polymerization of hydrophilic functional monomers.⁹⁷ The introduction of hydrophilic polymer layers onto the MIP microspheres significantly improved their surface hydrophilicity and suppressed the hydrophobically driven nonspecific interactions between the MIPs and template molecules, thus leading to MIPs with water-compatible template binding properties. Other approaches include preparation of MIP microspheres with surface-grafted hydrophilic polymer brushes by the facile one-pot RAFTPP mediated by hydrophilic macromolecular chain-transfer agents and combined use of RAFTPP and successive surface-initiated RAFT polymerization.^98,99 The use of CRPP methods is continuously blooming in the area of molecular imprinting.^102–105

5. Other advances in MIPs-PP

There are many appealing features of MIPs-PP which adds benefits of sensitivity and selectivity to the analytical method, meanwhile there are several advances blooming in the area. The synthesis of MIPs-PP is combined with other approaches to enhance the properties of resulting material. Some recent advances in the field are discussed in this section.

Dummy template approach is a use of a structurally similar compound as a substitute for the template compound. This approach is often used in molecular imprinting if the template is too toxic to handle or not available in sufficient amount. Moreover, this is also used to circumvent the problems of template bleeding in SPE.¹⁰⁶ Dummy template approach was to prepare the group selective MIPs-PP by Wu et al.¹⁰⁷ who prepared MIPs-PP using danthron as dummy template with MAA and EGDMA in a mixture of 75 : 25 acetone–toluene. The resulting polydispersed MIPs-PP revealed group selectivity for anthraquinones and six anthraquinones were detected in MISPE of slimming tea sample. The limit of quantification was in the range of mg kg⁻¹ and rapid separation of anthraquinones was observed. In other work, MIPs-PP was prepared for the simultaneous isolation and determination of five phthalate esters in plastic bottled beverages with the aim to avoid the template bleeding.¹⁰⁸ The diisononyl phthalate was used as dummy template with AM and DVB in toluene and acetonitrile and the MIPs-PP showed group selectivity and affinity for the five phthalate esters in the plastic bottled functional beverages. The average recoveries of the five phthalate esters in spiked samples ranged from 84 to 96% and the effect of template leakage was eliminated resulting in the successful determination of phthalate esters in complicated matrix.

Ultrasonication of pre-polymerization mixture was performed to enhance the rate of polymerization and to achieve homogeneous chain growth, greater yields, and milder conditions during the process of polymerization.^109,110 MAA and EGDMA were polymerized in acetonitrile for the template caffeic acid. The resulting MIPs microspheres exhibited narrow size distribution of particles with excellent yields and it was observed that the MIPs-PP prepared at comparatively lower temperature of 40 °C exhibited the best binding performance with faster binding kinetics.

Magnetic nanoparticles as sorbents have received considerable attention in sample pretreatment owing to its several advantages over traditional SPE.² A new hybrid magnetic material, where macrocyclic host molecule, pillar[5]arene, was functionalized on Fe₃O₄ magnetic nanoparticles via covalent bonds. This material was successfully utilized as magnetic SPE sorbent for the determination of trace pesticides in beverage which resulted in development of the MIPs-PP with fast binding kinetics of analyte.¹¹¹ MIPs-PP were developed for the determination of trace pesticides in beverage samples and the effects of various parameters, i.e., the amount of adsorbents, extraction and desorption times, desorption solvent, ionic strength and sample pH were optimized. The detection limits for seven pesticides were found in the ng mL⁻¹ range.

Magnetic MIPs were prepared using Fe₃O₄ nanoparticles and used for selective adsorption of 3-methylindole from fuel.¹¹² Synthesis of magnetic nanoparticles using precipitation polymerization is not a true precipitation polymerization since it involves Fe₃O₄ magnetic nanoparticles in the prepolymerization mixture. In the synthesis procedure, Fe₃O₄ was synthesized by coprecipitation method and was modified by MAA. Then, polymers were anchored on the surface of modified magnetic nanoparticles. The morphology of resultant MIPs was irregular and in the form of aggregate. However, these magnetic MIPs-PP exhibited high good recognition ability and fast binding kinetics for the target 3-methylindole.

A variant to the one step precipitation polymerization is development of two step core–shell MIPs which involves preparation of polymer microspheres core particles by precipitation polymerization of crosslinker in the first step. Thereafter, the core particles are used as seed particles in the synthesis of molecularly imprinted core–shell particles by copolymerization of monomer and crosslinker in presence of the template molecules.¹¹³ Thiabendazole, a fungicide was imprinted utilizing this concept for its detection in citrus fruits and orange juice samples.¹¹⁴ Poly-DVB core particles were used as seed particles in the production of MIP shells by copolymerization of DVB with MAA in the presence of thiabendazole in acetonitrile. The morphology assessment of core–shell MIPs by SEM and TEM showed formation of microspores around 4.0 μm with narrow size distribution and smooth surfaces. The MISPE resulted in high recovery of thiabendazole without including laborious sample clean-up step in short time span.

Core–shell MIPs were also prepared for bisphenol A, a MIP core around 3 μm in size was prepared by precipitation polymerization.¹¹⁵ These cores were further used in preparation of shell by optimization of solvent. The core–shell MIP revealed good selectivity and binding capacity toward bisphenol A in MISPE. Moreover, leakage of bisphenol A was not observed in extraction procedure which suggested potential application of MIPs-PP in trace analysis area.

Li et al.¹¹⁶ developed the MIP-coated magnetic nanoparticles by precipitation polymerization using TFMAA and EGDMA as functional monomer and crosslinker respectively. The morphology of MIPs was irregular. The MIPs sorbent reached the equilibrium quickly due to the presence of selective binding sites at the surface of MIP-coated magnetic nanoparticles and exhibited large adsorption capacity and 43.46 nmol g⁻¹ tadalafil was detected from the herbal sexual health product. Moreover, MIP-coated magnetic nanoparticles could be separated with an external magnetic field.

6. Concluding remarks

The MIPs prepared using precipitation polymerization (MIPs-PP) not only exhibits beaded and particulate morphology in micro to nano range but also selectivity for the target material. Moreover, it is possible to tune the particle size of MIPs-PPs. The performance of MIPs-PP is superior as compared to the conventional sorbent used in the chromatography. This is evident from the literature dealing with the application of MIPs-PP for extraction of target molecules from different complex matrices including environmental, biological and food. MIPs-PP can be applied in conjugation with other analytical technique. However, influence of template on morphology, large amount of solvent, longer polymerization time and attainment of desired morphology are some issues which needs to be addressed. Controlled living radical precipitation polymerization method can provide controlled preparation of MIPs with tailor-made structures, superior water compatibility and improved binding properties. MIPs-PP can be used successfully to bind variety of target molecules from various complex matrices.

Abbreviations

AA	Acrylic acid
AM	Acrylamide
DFT	Density functional theory
DVB	Divinyl benzene
EGDMA	Ethylene glycol dimethacrylate
HEMA	Hydroxyl ethyl methacrylate
HPLC	High performance liquid chromatography
MAA	Methacrylic acid
MBAA	N,N-Methylenebisacrylamide
MIPs	Molecularly imprinted polymers
MIPs-PP	MIPs produced using precipitation polymerization
MISPE	Molecularly imprinted solid phase extraction
SPE	Solid-phase extraction
THF	Tetrahydrofuran
TFMAA	Trifluoromethacrylic acid
TRIM	Trimethylol propane trimethacrylate
2-VP	2-Vinylpyridine
4-VP	4-Vinyl pyridine

References

1. K. Haupt, Molecular Imprinting, Springer-Verlag, Berlin Heidelberg, 2012, vol. 325, pp. 1–28.
2. L. Chen and B. Li, Anal. Methods, 2012, 4, 2613–2621.
3. G. Vasapollo, R. Del Sole, L. Mergola, M. R. Lazzoi, A. Scardino, S. Scorrano and G. Mele, Int. J. Mol. Sci., 2011, 12, 5908–5945.
4. F. Liu, S. Zhang, G. Wang, J. Zhao and Z. Guo, RSC Adv., 2015, 5, 22811–22817.
5. J. Wang, P. A. G. Cormack, D. C. Sherrington and E. Khoshdel, Angew. Chem., Int. Ed. Engl., 2003, 42, 5336–5338.
6. J. Dai, Y. Zhang, M. Pan, L. Kong and S. Wang, J. Agric. Food Chem., 2014, 62, 5269–5274.
7. J. K. Oh, R. Drumright, D. J. Siegwart and K. Matyjaszewski, Prog. Polym. Sci., 2008, 33, 448–477.
8. S. Muratsugu and M. Tada, Acc. Chem. Res., 2013, 46, 300–311.
9. P. Regal, M. Díaz-Bao, R. Barreiro, A. Cepeda and C. Fente, Cent. Eur. J. Chem., 2012, 10, 766–784.
10. C. He, Y. Long, J. Pan, K. Li and F. Liu, J. Biochem. Biophys. Methods, 2007, 70, 133–150.
11. B. Sellergren, Molecularly Imprinted Polymers: Man-Made Mimics of Antibodies and their Application in Analytical Chemistry, Elsevier Science, Netherlands, 2003, vol. 1, p. 309.
12. Z. Luo, A. Zeng, P. Zheng, P. Guo, W. Du, K. Du and Q. Fu, Anal. Methods, 2014, 6, 7865–7874.
13. E. Zambrzycka, U. Kiedysz, A. Z. Wilczewska, B. Lesniewska and B. Godlewska, Anal. Methods, 2013, 5, 3096–3105.
14. S. A. Mohajeri, G. Karimi, J. Aghamohammadian and M. R. Khansari, J. Appl. Polym. Sci., 2011, 121, 3590–3595.
15. L. Ye, P. A. G. Cormack and K. Mosbach, Anal. Chim. Acta, 2001, 435, 187–196.
16. C. Cacho, E. Turiel, A. Martin-Esteban, C. Pérez-Conde and C. Cámara, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci., 2004, 802, 347–353.
17. K. Yoshimatsu, K. Reimhult, A. Krozer, K. Mosbach, K. Sode and L. Ye, Anal. Chim. Acta, 2007, 584, 112–121.
18. Y. Wang, Y. Ding, F. Rong and D. Fu, Polym. Bull., 2012, 68, 1255–1270.
19. J. Lai, M. Yang, R. Niessner and D. Knopp, Anal. Bioanal. Chem., 2007, 389, 405–412.
20. J. Wang, P. A. G. Cormack, D. C. Sherrington and E. Khoshdel, Pure Appl. Chem., 2007, 79, 1505–1519.
21. J. Wackerlig and P. Lieberzeit, Sens. Actuators, B, 2015, 207, 144–157.
22. H. Jiang, H. Chen, Y. Liang and X. Liu, Polym. Adv. Technol., 2011, 22, 1555–1562.
23. E. C. Goh and H. D. Stover, Macromolecules, 2002, 35, 9983–9989.
24. R. Schirhagl, Anal. Chem., 2014, 86, 250–261.
25. L. Chen, S. Xu and J. Li, Chem. Soc. Rev., 2011, 40, 2922–2942.
26. A. Beltran, F. Borrull, P. A. G. Cormack and R. M. Marce, Trends Anal. Chem., 2010, 29, 954–965.
27. X. Shen, C. Xu and L. Ye, Ind. Eng. Chem. Res., 2013, 52, 13890–13899.
28. L. Figueiredo, G. L. Erny, L. Santos and A. Alves, Talanta, 2016, 146, 754–765.
29. S. Das, G. W. Brudvig and R. H. Crabtree, Chem. Commun., 2008, 4, 413–424.
30. X. Shen, L. Zhu, N. Wang, L. Ye and H. Tang, Chem. Commun., 2012, 48, 788–798.
31. W. Cheong, S. Yang and F. Ali, J. Sep. Sci., 2013, 36, 609–628.
32. A. McCluskey, C. Holdsworth and M. Bowyer, Org. Biomol. Chem., 2007, 5, 3233–3244.
33. L. Nováková and H. Vlcková, Anal. Chim. Acta, 2009, 656, 8–35.
34. M. Resmini, Anal. Bioanal. Chem., 2012, 402, 3021–3026.
35. M. Talha Gokmen and F. Du Prez, Prog. Polym. Sci., 2012, 37, 365–405.
36. K. Karim, F. Breton, R. Rouillon, E. V. Piletska, A. Guerreiro, I. Chianella and S. A. Piletsky, Adv. Drug Delivery Rev., 2005, 57, 1795–1808.
37. H. Yan and K. Row, Int. J. Mol. Sci., 2006, 7, 155–178.
38. Q. Yan, T. Zhao, Y. Bai, F. Zhang and W. Yang, J. Phys. Chem. B, 2009, 113, 3008–3014.
39. W. H. Li and H. D. H. Stöver, J. Polym. Sci., Part A: Polym. Chem., 1998, 36, 1543–1551.
40. F. Lime and K. Irgum, Macromolecules, 2009, 42, 4436–4442.
41. X. Shi, L. Xu, H. Duan, Y. Huang and Z. Liu, Electrophoresis, 2011, 32, 1348–1356.
42. M. Esfandyari-Manesha, M. Javanbakht, F. Atyabi, A. Mohammadi, S. Mohammadi, B. Akbari-Adergani and R. Dinarvand, Mater. Sci. Eng., C, 2011, 31, 1692–1699.
43. E. Asadi, S. Azodi-Deilami, M. Abdouss and S. Khaghani, Appl. Biochem. Biotechnol., 2012, 167, 2076–2087.
44. A. Abouzarzadeh, M. Forouzani, M. Jahanshahi and N. Bahramifar, J. Mol. Recognit., 2012, 25, 404–413.
45. Y. Shi, H. Lv, X. Lu, Y. Huang, Y. Zhang and W. Xue, J. Mater. Chem., 2012, 22, 3889–3898.
46. E. Benito-Peña, F. Navarro-Villoslada, S. Carrasco, S. Jockusch, M. Ottaviani and M. Moreno-Bondi, ACS Appl. Mater. Interfaces, 2015, 7, 10966–10976.
47. S. Noee, N. Salimraftar, M. Abdouss and G. Riazi, Polym. Int., 2013, 62, 1711–1716.
48. Y. Liu, K. Hoshina and J. Haginaka, Talanta, 2010, 80, 1713–1718.
49. N. Pérez-Moral and A. G. Mayes, Anal. Chim. Acta, 2004, 504, 15–21.
50. W. Zhang, N. Tan, X. Jia, G. Wang, W. Long, X. Li, S. Liao and D. Hou, Mater. Sci. Eng., 2015, 53, 166–174.
51. A. Chauhan, T. Bhatia, M. Gupta, P. Pandey, V. Pandey, P. Saxena and M. K. R. Mudiam, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci., 2015, 1001, 66–74.
52. S. Chaitidou, O. Kotrotsiou, K. Kotti, O. Kammona, M. Bukhari and C. Kiparissides, Mater. Sci. Eng., B, 2008, 152, 55–59.
53. H. Ebrahimzadeh, K. Molaei, A. A. Asgharinezhad, N. Shekari, Z. DehghaniAntonelle Pardo, L. Mespouille, P. Dubois, P. Duez and B. Blankert, Anal. Chim. Acta, 2013, 767, 155–162.
54. A. Pardo, L. Mespouille, P. Dubois, P. Duez and B. Blankert, Cent. Eur. J. Chem., 2012, 3, 751–765.
55. X. Yina, Q. Liub, Y. Jiang and Y. Luo, Spectrochim. Acta, Part A, 2011, 79, 191–196.
56. F. Chen, G. Wang and Y. Shi, J. Sep. Sci., 2011, 34, 2602–2610.
57. Á. Navarro, M. Romeroa, J. F. Sánchez, P. A. G. Cormack, A. Carretero and A. Fernández-Gutiérrez, J. Chromatogr. A, 2011, 1218, 7289–7296.
58. S. Pardeshi, R. Dhodapkar and A. Kumar, Food Chem., 2014, 146, 385–393.
59. N. Funaya and J. Haginaka, J. Chromatogr. A, 2012, 1248, 18–23.
60. C. Miura, H. Li, H. Matsunaga and J. Haginaka, J. Pharm. Biomed. Anal., 2015, 114, 139–144.
61. H. Zhu, Y. Wang, Y. Yuan and H. Zeng, Anal. Methods, 2011, 3, 348–355.
62. Y. Yuan, Y. Wang, M. Huang, R. Xu, H. Zeng, C. Nie and J. Kong, Anal. Chim. Acta, 2011, 695, 63–72.
63. J. Fan, Z. Tian, S. Tong, X. Zhang, Y. Xie, R. Xu, Y. Qin, L. Li, J. Zhu and X. Ouyang, Food Chem., 2013, 141, 3578–3585.
64. W. Zhanga, H. Zhang, Q. Zhang, Y. Cui, Z. Wu, R. Zheng and L. Liu, Sep. Purif. Technol., 2011, 81, 411–417.
65. T. de Boer, R. Mol, R. A. de Zeeuw, G. J. de Jong, D. C. Sherrington, P. A. G. Cormack and K. Ensing, Electrophoresis, 2002, 23, 1296–1300.
66. A. Beltran, F. Borrull, P. A. G. Cormack and R. M. Marcé, J. Chromatogr. A, 2011, 1218, 4612–4618.
67. J. Lai, L. Xie, H. Sun and F. Chen, Anal. Bioanal. Chem., 2013, 405, 4269–4275.
68. Z. Yu, Q. Su, Y. Tang and Z. Xu, J. Appl. Polym. Sci., 2013, 128, 3425–3431.
69. Y. Liu, G. Zhang, L. Deng, J. Lei, L. Wang and J. He, Anal. Methods, 2014, 6, 684–689.
70. Y. Wang, Q. Liu, F. Rong and D. Fu, Appl. Surf. Sci., 2011, 257, 6704–6710.
71. A. Beltran, F. Borrull, P. A. G. Cormack and R. M. Marcé, J. Chromatogr. A, 2011, 1218, 4612–4618.
72. S. Pardeshi, R. Dhodapkar and A. Kumar, J. Mol. Model., 2012, 18, 4797–4810.
73. M. B. Gholivand and M. Khodadadian, Talanta, 2011, 85, 1680–1688.
74. L. Jun-Bo, S. Yang, T. Shan-Shan and J. Rui-Fa, J. Sep. Sci., 2015, 38, 1065–1071.
75. X. Shen, L. Zhu, N. Wang, L. Ye and H. Tang, Chem. Commun., 2012, 48, 788–798.
76. V. Pichon and F. Chapuis-Hugon, Anal. Chim. Acta, 2008, 622, 48–61.
77. C. Dai, X. Zhou, Y. Zhang, S. Liu and J. Zhang, J. Hazard. Mater., 2011, 198, 175–181.
78. C. Dai, S. Geissen, Y. Zhang, Y. Zhang and X. Zhou, Environ. Pollut., 2011, 159, 1660–1666.
79. E. Rodríguez, F. Navarro-Villoslada, E. Benito-Pena, M. Marazuela and M. Moreno-Bondi, Anal. Chem., 2011, 83, 2046–2055.
80. P. Xiao, Y. Dudal, P. Corvini, P. Spahr and P. Shahgaldian, React. Funct. Polym., 2012, 72, 287–293.
81. P. Narula, V. Kaur, R. Singh and S. Kansal, J. Sep. Sci., 2014, 37, 3330–3338.
82. A. Beltran, R. M. Marcé, P. A. G. Cormack and F. Borrull, Anal. Chim. Acta, 2010, 677, 72–78.
83. T. Jing, X. Gao, P. Wang, Y. Wang, Y. Lin, X. Hu, Q. Hao, Y. Zhou and S. Mei, Anal. Bioanal. Chem., 2009, 393, 2009–2018.
84. Z. Qiujin, W. Liping, W. Shengfang, J. Wasswa, G. Xiaohong and T. Jian, Food Chem., 2009, 113, 608–615.
85. X. Xu, S. Liang, X. Meng, M. Zhang, Y. Chen, D. Zhao and Y. Li, J. Chromatogr. B: Anal. Technol. Biomed. Life Sci., 2015, 988, 182–186.
86. J. Du, R. Gao, H. Yu, X. Li and H. Mu, J. Pharm. Anal., 2014, 5, 200–206.
87. L. Gao, X. Li, Q. Zhang, J. Dai, X. Wei, Z. Song, Y. Yan and C. Li, Food Chem., 2014, 156, 1–6.
88. J. He, Y. Shen, S. Chen, H. Wei, J. Zhu, L. You and K. Lu, J. Sep. Sci., 2011, 34, 2739–2744.
89. H. Yan, M. Gao and J. Qiao, J. Agric. Food Chem., 2012, 60, 6907–6912.
90. H. Cao, F. Xu, D. Li, X. Zhang and J. Yu, Res. Chem. Intermed., 2013, 39, 2321–2337.
91. M. Behbahani, P. Hassanlou, M. M. Amini, H. Moazami, H. Abandansari, A. Bagheri and S. Zadeh, Food Analytical Methods, 2015, 8, 558–568.
92. K. Yang, M. Berg, C. Zhao and L. Ye, Macromolecules, 2009, 42, 8739–8746.
93. H. Zang, Eur. Polym. J., 2013, 49, 579–600.
94. B. Y. Zu, G. Q. Pan, X. Z. Guo, Y. Zhang and H. Q. Zhang, J. Polym. Sci., Part A: Polym. Chem., 2009, 47, 3257–3270.
95. J. S. Jiang, Y. Zhang, X. Z. Guo and H. Q. Zhang, RSC Adv., 2012, 2, 5651–5662.
96. J. Y. Li, B. Y. Zu, Y. Zhang, X. Z. Guo and H. Q. Zhang, J. Polym. Sci., Part A: Polym. Chem., 2010, 48, 3217–3228.
97. G. Q. Pan, Y. Zhang, X. Z. Guo, C. X. Li and H. Q. Zhang, Biosens. Bioelectron., 2010, 26, 976–982.
98. G. Q. Pan, Y. Zhang, Y. Ma, C. X. Li and H. Q. Zhang, Angew. Chem., Int. Ed., 2011, 50, 11731–11734.
99. Y. Ma, Y. Zhang, M. Zhao, X. Z. Guo and H. Q. Zhang, Chem. Commun., 2012, 48, 6217–6219.
100. G. Q. Pan, B. Y. Zu, X. Z. Guo, Y. Zhang, C. X. Li and H. Q. Zhang, Polymer, 2009, 50, 2819–2825.
101. S. Xu, J. Li and L. Chen, Talanta, 2011, 85, 282–289.
102. L. Zhao, F. Zhao and B. Zeng, Biosens. Bioelectron., 2014, 60, 71–76.
103. H. Zhang, Polymer, 2014, 55, 699–714.
104. H. Wang, X. Dong and M. Yang, Trends Anal. Chem., 2012, 31, 96–108.
105. J. Dai, Y. Zou, Z. Zhou, X. Dai, J. Pan, P. Yu, T. Zou, Y. Yan and C. Li, RSC Adv., 2014, 4, 1965–1973.
106. J. Yin, Z. Meng, Y. Zhu, M. Song and H. Wang, Anal. Methods, 2011, 3, 173.
107. X. Wu, S. Liang, X. Ge, Y. Lv and H. Sun, J. Sep. Sci., 2015, 38, 1263–1270.
108. X. Cheng and G. Yang, J. Agric. Food Chem., 2012, 60, 5524–5531.
109. N. Phutthawong and M. Pattarawarapan, Polym. Bull., 2013, 70, 691–705.
110. N. Phutthawong and M. Pattarawarapan, J. Appl. Polym. Sci., 2013, 128, 3893–3899.
111. M. Tian, D. Chen, Y. Sun, Y. Yang and Q. Jia, RSC Adv., 2013, 3, 22111–22119.
112. D. Niu, Z. Zhou, W. Yang, Y. Li, L. Xia, B. Jiang, W. Xu, W. Huang and T. Zhu, J. Appl. Polym. Sci., 2013, 130, 2859–2866.
113. N. Marchyk, J. Maximilien, S. Beyazit, K. Haupt and B. Bui, Nanoscale, 2014, 6, 2872–2878.
114. F. Barahona, E. Turiel, P. A. G. Cormack and A. Martin Esteban, J. Sep. Sci., 2011, 34, 217–224.
115. Y. Wang, Q. Liu, F. Rong and D. Fu, Appl. Surf. Sci., 2011, 257, 6704–6710.
116. Y. Li, M. Ding, S. Wang, R. Wang, X. Wu, T. Wen, L. Yuan, P. Dai, Y. Lin and X. Zhou, ACS Appl. Mater. Interfaces, 2011, 3, 3308–3315.

---