Molecularly imprinted polymer-hollow fiber microextraction of hydrophilic fluoroquinolone antibiotics in environmental waters and urine samples

Journal of Chromatography A, 1587 (2019) 42–49

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Journal of Chromatography A journal homepage: www.elsevier.com/locate/chroma

Molecularly imprinted polymer-hollow fiber microextraction of hydrophilic fluoroquinolone antibiotics in environmental waters and urine samples Francisco Barahona, Beatriz Albero, José Luis Tadeo, Antonio Martín-Esteban ∗ Departamento de Medio Ambiente, INIA, Carretera de A Coru˜ na km 7, 28040, Madrid, Spain

a r t i c l e

i n f o

Article history: Received 28 August 2018 Received in revised form 4 December 2018 Accepted 9 December 2018 Available online 10 December 2018 Keywords: Fluoroquinolones Molecularly imprinted polymers MIP hollow fiber Microextraction Aqueous samples Miniaturized extraction techniques

a b s t r a c t In the present work, the preparation of a new selective molecularly imprinted polymer (MIP) for the family of fluoroquinolones (FQs) in the pores of polypropylene hollow fibers (HFs) is proposed. The resulting MIP-HFs, which combine solid-phase microextraction (SPME) and molecular imprinting technologies, were used to develop a selective microextraction methodology (MIP-HFM) to determine selected FQs danofloxacin, norfloxacin, enrofloxacin and ciprofloxacin in real samples of environmental and biological interest. Measurements during the optimization of the MIP-HFM and its application to the analyses of real samples were performed by HPLC-UV and HPLC-MS/MS. In order to establish optimum rebinding conditions, the effect of key experimental parameters such as loading media, extraction time and stirring-rate were studied. The developed MIP composites exhibited recognition properties towards the selected hydrophilic antibiotics in non-polar media (toluene) and in polar protic systems such as methanol and methanol/water solutions, up to 20% water content. Recoveries by the developed method for all FQs tested in surface water, groundwater and urine spiked with the analytes of interest at two different concentration levels were within 9.4–24.5 %, with a relative standard deviation, generally <20% (n = 3). The detection limits were within 0.1–10 ␮g L−1 , depending upon the antibiotic and the type of sample. © 2018 Elsevier B.V. All rights reserved.

1. Introduction Since its introduction in 1990 by Arthur and Pawliszyn, solid-phase microextraction (SPME) has attracted the interest of analytical chemists as a useful microextraction technique and it is currently a preferred technique in many analytical tests [1]. The advantages of SPME are common to any other miniaturized techniques: the reduction or absence of organic solvents; shorter extraction times and possibility of simultaneous processing of several samples; smaller size of the extraction systems that facilitates its implementation in portable devices and field work; enhanced sensitivity due to concentration of the analytes in a small extractant phase; etc [2]. A proper selection of the SPME sorbent is a key factor in the success of the analysis [3,4]. In general, the polarity of the fiber should be as similar as possible to that of the analyte of interest. In this sense, nowadays a great variety of fibers commercially

∗ Corresponding author. E-mail address: [email protected] (A. Martín-Esteban). https://doi.org/10.1016/j.chroma.2018.12.015 0021-9673/© 2018 Elsevier B.V. All rights reserved.

available covers a wide range of polarities (i.e. Carbowax/DVB for polar compounds or polydimethylsiloxane for hydrophobic compounds). Also, both the fiber thickness and the porosity of the sorbent will influence the final extraction efficiency. Besides, other physical and chemical parameters such as temperature, exposition time, agitation, pH, or ionic strength (“salting out” effect) of the sample can be also optimised. Molecularly imprinted polymers (MIPs) are synthetic materials that exhibit selective recognition towards the target molecule for which they were prepared. MIPs are synthesized by polymerizing functional and cross-linking monomers around a template molecule, which afterwards will be extracted generating binding sites with shape, size and functionalities complementary to the target analyte. Their use as selective sorbents in solid-phase extraction, so-called molecularly imprinted solid-phase extraction (MI-SPE), is by far the most advanced technical application of MIPs [5–7]. MIPs provide clean extracts and prevent the introduction of undesired compounds in the detection systems that can hinder the determination of the analytes and potentially damage the detection equipment. Because of these abilities, MIPs are consid-

F. Barahona et al. / J. Chromatogr. A 1587 (2019) 42–49

ered a very useful tool for analytical chemists [8]. Similarly, SPME has also taken advantage of the selectivity provided by MIPs to the extraction techniques. The number of applications combining MIPs and SPME (MI-SPME) in a variety of formats confirms such a synergy [9,10]. Works using MI-SPME make use of different strategies for the preparation of ␮-sized selective sorbents and subsequently they propose different modes of use and applications such as molecularly imprinted magnetic solid-phase extraction [11]; in tube MI-SPME [12]; molecularly imprinted polymer stir bar sorptive extraction [13–15] or the use of MIP fibers, among others. MIP fibers can be a composite formed by a support coated by MIP [16,17], or be formed by a MIP monolith [18,19]. Recently, the modification of commercial polypropylene microfiltration capillaries has been proposed to generate MIP-hollow fibers (MIP-HFs) for diethylstilbestrol [20], organochlorine pesticides [21], estrogens [22] and triazines [23], according to different polymerization strategies. Fluoroquinolones (FQs) are an important group of antibiotics widely used in human and veterinary medicine. FQs are 6-fluorinated piperazinyl derivatives of nalidixic acid, the first commercial quinolone. Originally applied in the treatment of urinary tract infections, FQs are nowadays widely used in the treatment of respiratory diseases and bacterial infections in humans. Likewise, some of them have been approved for use in farm animals intended for human consumption, such as cattle, pigs, poultry as well as in aquaculture [24,25]. Because such a widespread usage, FQs are considered as an emergent class of environmental pollutants [26], and they have attracted the attention of scientists to develop new analytical methods to extract and determine their presence in the environment [27,28], in biological fluids [29] and food [30]. In the present work, the preparation of a new selective MIP for the family of FQs in the pores of polypropylene tubular membranes is proposed. Given the marked hydrophilicity of these compounds, the prepared MIP-HFs must demonstrate selective recognition towards the target analytes in a suitable polar medium. The MIP-HFs will be applied to the development of a microextraction methodology to determine selected FQs in real aqueous samples of environmental and biological interest. 2. Materials and methods 2.1. Chemicals and reagents Norfloxacin, (NOR, purity ≥ 98.0%), enrofloxacin (ENRO, purity ≥ 99.8%), ciprofloxacin (CIP, purity ≥ 99.0%) and danofloxacin (DAN, purity ≥ 99.6%) were obtained from Sigma-Aldrich (St Louis, MO, USA). Stock standard solutions (1 g L−1 ) were prepared in acetonitrile containing 2% ammonia and stored at −18 ◦ C. Methacrylic acid (MAA), ethylene glycol dimethacrylate (EGDMA), and 2,2 azobisisobutyronitrile (AIBN) were purchased from Sigma-Aldrich. EGDMA and MAA were freed from stabilizers by distillation under reduced pressure, and AIBN was recrystallised from methanol prior to use. All organic solvents and purified water were HPLC-grade and purchased from Scharlab (Barcelona, Spain). All other chemicals used were of analytical reagent-grade and obtained from Panreac Quimica (Barcelona, Spain)

43

segments were sealed by applying mechanical pressure. A scheme of the polymerization procedure for the preparation of MIP-HFs is displayed in Fig. 1. A polymerization mixture was prepared containing the template molecule (ENRO, 0.3 mmol), functional monomer (MAA, 1.2 mmol), cross-linker (EGDMA, 6.0 mmol), initiator (AIBN, 3.9 mmol), and porogen (toluene, 1.73 mL). A number of 15 segments of HF were placed inside a glass Pasteur pipette which acted as reactor for the polymerization reaction. The pipette containing the pieces of HF was then filled with the polymerization solution. One piece of rubber and one of Teflon sealed both the lower and upper ends of the pipette, respectively. The polymerization reaction takes place in two steps: 1) the pipette containing the HFs immersed in the polymerization mixture is placed into a temperature controllable incubator (S160D mode, Barloworld Scientific, Staffordshire, UK) with the temperature set to 60 ◦ C; 2) after a determined time, the so-called MIP-HFs are removed from the reaction mixture, separated each other and placed into an empty 15 mL vial inside the incubator at 60 ◦ C for further 60 min. Finally, MIP-HFs are immersed in a methanol: acetic acid (1:1, v/v) solution for 2 h, under vigorous stirring in order to remove the template. The absence of leaching was verified by a flat baseline in HPLC-UV analysis. Non-imprinted polymer materials (NIP-HF) were also prepared as described above but without the addition of the template. 2.3. Materials characterization Scanning Electron Microscopy (SEM) micrographs were obtained at the Centro de Microscopıa Electronica Luis Bru, Universidad Complutense de Madrid, using a JEOL JSM-6335 F (JEOL, Tokyo, Japan). Synthesized materials were characterized by IR spectroscopy on a Jasco FTIR-460 Plus spectrophotometer. All spectra were recorded by attenuated total reflectance between 4000 and 500 cm−1 , with the sample in the solid state without other treatment. 2.4. Molecularly imprinted polymer- hollow fiber microextraction (MIP-HFM) The MIP-HFs were conditioned during 5 min by immersion in the same solvent as the sample medium. A 0.8 mL volume of aqueous sample to be extracted was mixed with methanol 1:5 (v/v) up to a volume of 4 mL in a screw cap 4.5 mL glass vial. The molecularly imprinted polymer-coated hollow fiber microextraction (MIP-HFM) process was performed by direct immersion of the conditioned fibers in the sample for 30 min, subjected to orbital stirring using Vibramax 100 agitator (Heidolph, Kelheim, Germany) at controlled speed rate (at 750 rpm). Then, MIP-HFs were washed in 1.8 mL clean methanol for 10 min under stirring (750 rpm) in order to remove nonspecific interactions. Finally, the target analytes were desorbed by agitation at 750 rpm for 15 min using 400 ␮L of 2% formic acid solution in methanol inside 0.4 mL vial inserts. The extracts were diluted 1:1 (v/v) with water prior to HPLC analysis. Between extractions, MIP-HFs were reconditioned by immersion in methanol for 15 min.

2.2. Preparation of polymers 2.5. HPLC-UV The polymerization procedure was adapted to that described in [23] by the authors. For a better clarity, it is briefly explained below. Commercial porous Q3/2 polypropylene hollow fiber (PP HF) from Membrana (Wupertal, Germany) with an internal diameter of 600 ␮m, a 200 ␮m of wall thickness and 0.2 ␮m pores were cut in pieces of 6 cm length and carefully weighed. The ends of these

During optimization, the measurements were carried out by means of an 1100 LC instrument equipped with a gradient pump, autosampler, and programmable UV–vis detector from Agilent Technologies (Wilmington, DE, USA). The analytes were separated on an Atlantis dC18 HPLC column (150 × 3.9 mm, 3 ␮m) from

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F. Barahona et al. / J. Chromatogr. A 1587 (2019) 42–49

Fig. 1. Scheme of the 2-step polymerization procedure for the preparation of MIP-HFs.

Waters (Massachusetts, USA). The mobile phase was a linear gradient prepared from diluted formic acid, pH 2.5, 3.16 mmol L−1 (component A) and acetonitrile (component B); the program was from 96% A-4% B to 50% A-50% B in 17 min, returning to the initial conditions in 3 min. The flow rate was set at 0.7 mL min−1 and the column temperature was controlled at 25 ◦ C. The UV detector was programmed to monitor at 280 nm. The injection volume was 50 ␮L.

3. Results and discussion 3.1. Polymerization and characterization

(1)

3.1.1. Synthesis The polymerization procedure described in the Materials and methods section allows the crosslinking of the polymer matrix inside the pores of hollow polypropylene fibers. First, for an efficient polymerization and subsequent molecular recognition, all components of the polymerization mixture must be appropriately dissolved in the polymerization mixture. Therefore, ENRO was chosen as template because of a greater solubility in the selected porogen, toluene. This solvent was chosen because it promotes the generation of polymers with a greater porosity compared to other organic solvents such as acetonitrile or methanol. The hydrophobic nature of the support (polypropylene) favors the absorption of the polymerization mixture (prepared in toluene) by the walls of the fibers. By sealing both ends of the fibers, the polymerization mixture cannot freely access the interior of the fiber cavity and soaks it from the outside. The mixture is retained in the pores of the walls by surface tension, hence restricting the polymerization also to the walls, keeping the lumen of the hollow fibers empty. In the case of not sealing the ends of the polyethylene fibers, the polymerization mixture penetrates inside the fibers by capillarity, filling the lumen, and the polymerization leads to polymer monoliths therein. Up to 15 fibers can be hosted inside the Pasteur pipette acting as polymerization reactor. The polymerization occurs at 60 ◦ C inside a preheated thermostatic incubator and it is divided in two parts. Firstly, the reaction initiates thermally and the liquid mixture gradually changes to gel state. At this stage, the polymer is sufficiently cross-linked inside the macropores of the PP fibers but these can still be separated from the polymerization medium. Based on preliminary studies, 32.5 min is chosen as an optimum reaction time, because longer polymerization times led to a solid block and shorter times led to a smaller amount of immobilized polymer in the walls of the fiber and to greater variability. A second step in the preparation of the polymer-HF composites takes place during one additional hour in the incubator set at 60 ◦ C in order to continue the polymerization of the gel mixture previously immobilized.

where na initial and na final are the number of moles of analyte originally present in the sample subjected to extraction and the number of moles of analyte finally present in the final solution after the complete extraction process, respectively; Va is the volume of solution prior to HPLC analysis; Vs is the volume of sample to be extracted; ca final is the concentration of analyte in the final solution; ca initial is the original analyte concentration within the sample. In case of a previous dilution of the original sample (Vo ), Vs is corrected using an appropriate dilution factor (df = Vo /Vs ).

3.1.2. Characterization of the MIP-HFs To assess the efficiency of the polymerization procedure, the fibers were subjected to gravimetrical analysis before and after modification. Table 1 includes the increment of mass attributed to the new polymer for MIP and NIP fibers of the same polymerization batch (5.0 and 4.5 mg, respectively). The RSD of the fibers that can be prepared in one single batch were lower than 13%. To ensure a satisfactory repeatability of the polymerization process,

2.6. HPLC-MS/MS Real samples were analyzed by LC–MS/MS with an Agilent 1200 liquid chromatograph coupled to an Agilent 6410 (Waldbronn, Germany) triple quadrupole mass spectrometer equipped with an electrospray ionization (ESI) source. The optimized ESI parameters were the following: drying gas temperature of 300 ◦ C, drying gas flow rate of 9 L min−1 ; nebulizer gas pressure of 45 psi and capillary voltage of 4000 V. The chromatographic separation was carried out using a Kinetex XB-C18 (100 mm x 3 mm i.d., 2.6 ␮m particle size) analytical column with a C18 security guard cartridge from Phenomenex (Torrance, CA, USA). LC separation of the analytes was performed injecting 10 ␮L with a gradient elution program at a flow rate set at 0.3 mL min−1 and a column temperature of 30 ◦ C. The mobile phases were acetonitrile as mobile phase A and 0.1% formic acid in water as mobile phase B with the following gradient program (with respect to phase B): 0 min, 95% B; 2 min, 80% B; 9 min, 40% B; 10 min, 5% B; 13 min, 5% B; 13.1 min, 95% B; 16 min, 95% B. A post-run time of 4 min was allowed before the next injection. Analyses were performed in the Multiple Reaction Monitoring (MRM) mode. The precursor and product ions used for quantification and confirmation of the studied FQs are shown in the supplementary material (table S1). 2.7. Recovery calculations Recovery (R, %) was calculated according to the following equation: R(%) = (nafinal /nainitial )x100 = [(Va /Vs )(cafinal /cainitial )]x100

F. Barahona et al. / J. Chromatogr. A 1587 (2019) 42–49 Table 1 Efficiency of the polymerization on NIP and MIP-HFs. Data represent the mass increase after polymerization. RSD (%) intra-batch corresponds to the measurement of 10 fibers. RSD (%) inter-batch was calculated with the measurements of three different batches of NIP and MIP-HFs. MIP

NIP a

m average (mg) RSD (%) a b

b

45

ing bands, respectively. The infrared spectra of MIP-HF presents additionally the peak of C O at 1150 cm−1 , and the intense peak of C O group at 1723 cm−1 indicating the presence of polymer successfully formed on the pores of the HF. 3.2. Evaluation of the MIP-HF re-binding capability

Intra-batch

Inter-batch

Intra-batcha

Inter-batchb

5.0 12.8

4.9 10.5

4.5 11.2

4.4 11.4

N = 10. N = 3.

Fig. 2. Scanning electron micrographs of (A) wall surface of original polypropylene hollow fiber; (B) wall surface of MIP-HF after modification. Magnification x300 for A and B.

the polymerization mixture is preheated inside the incubator for 2 min, before the addition of the initiator. The polymerization process was repeated several times under identical conditions. The repeatability of the polymerization process, in terms of inter-batch RSD (N = 3) was acceptable, with a RSD value in any case lower than 11.4%. Fig. 2A shows a SEM image of the commercial PP HF before modification. After polymerization, the new polymer fills the macropores of the original HF, as shown in Fig. 2B. The surface of the MIP-HF is macroscopically smooth and homogeneous. The interior of the modified fibers remains empty and the presence of the polymer is limited to the walls of the fibers. FT-IR spectroscopy was used for the characterization of PP HF and MIP-HF, and their corresponding spectra are shown in Supplementary Fig. S-1. The infrared spectra of PP HF presents bands at 2925, 2916, 2865 and 2837 cm−1 , which might be attributed to the characteristic CH2 (asymmetric), CH, and CH2 (symmetric) stretch-

Once the MIP is crosslinked in the walls of the PP fiber, it is necessary to extract all the template molecule to generate free specific binding sites for FQs. One advantage of the polymerization in the walls of the PP fibers is the facilitated diffusion of the target analytes. Likewise, the extraction of the template does not require an exhaustive technique such as the traditional Soxhlet extraction. Instead, stirring in methanol: acetic mixture (1:1, v/v) for 2 h was sufficient to remove the template molecule. The chromatographic analysis of blank extractions ensured the non-existence of template leaching. The optimization process of the MIP-HFM procedure was carried out by evaluating the molecular recognition of CIP, to prevent in case of an eventual presence of ENRO residues. Moreover, CIP has an intermediate polarity in contrast with ENRO, so that a more representative behavior of the rest of the compounds under study can be expected. 3.2.1. Loading, washing and elution media The specific recognition of FQs by the prepared MIP fibers in different media was evaluated. The microextraction procedure includes fiber conditioning, sample loading, washing step and elution. Initially, four loading solvents were evaluated: water, as it is a typical extraction medium for FQs, which are polar antibiotics of environmental and biological interest; methanol, because it is a protic and polar organic solvent; acetonitrile, because it is an organic solvent of intermediate polarity; and toluene, as it is the polymerization medium. Before the loading step, a conditioning step by immersion in the same clean loading solvent for 5 min was performed for all the solvents. Subsequently, the fibers were immediately immersed in 4 mL spiked solutions of CIP at 0.5 mg L−1 inside a screw cap glass vial. After stirring at 600 rpm during 30 min, the MIP-HFs were washed in a 2 mL vial containing 1.8 mL of clean loading solvent for 10 min at 600 rpm. The washing step in any MIP extraction aims to remove eventual undesired substances and compounds non-specifically attached to the polymer while keeping most of the target analyte-MIP specific interactions established during the loading step unaltered. The elution was performed during 15 min at 750 rpm using 400 ␮L acid solution placed inside a 400 ␮L flat bottom glass insert. Both 50% acetic acid in methanol (v/v) and 2% formic acid in methanol (v/v) were able to elute all the analyte retained by the polymer. Between the two options, formic acid was preferred because of a better chromatographic performance, resulting in better-resolved and defined peaks than in the case of the elution with acetic acid. This phenomenon can be explained according to the compatibility with the HPLC mobile phase, which contains 0.1% formic acid. The recoveries obtained under these microextraction conditions are shown in Fig. 3. As it was expected, MIP-HFs showed the greatest recoveries when toluene was the loading solvent, with recovery values of 28.5 and 9.9% for MIP and NIP, respectively. These values demonstrate that an effective imprinting was achieved. However, given the polar nature of FQs it is unlikely to find them naturally in such a medium, neither as a given sample nor as extractant or clean-up medium. Thus, the recognition in more polar media was investigated. When methanol was the loading solvent, recoveries of CIP were 13.6 and 2.9% for MIP and NIP, respectively. This behavior is in accordance with previous studies in which the specific recognition of FQs in methanol by a different MIP is documented [28] [30]. In the case of water, the extraction was not efficient at all, which can be explained by the preferred affinity of the FQs to

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Fig. 3. Recoveries (%) obtained after extraction of 4 mL spiked solutions of CIP at 0.5 mg L−1 concentration level in different solvents, using MIP and NIP fibers.

3.3. Optimization of MIP-HFM procedure

Fig. 4. Extraction recovery of CIP in 4 mL samples at 0.5 mg L−1 concentration level and increasing water content in the loading solvent composition.

the aqueous medium instead of the MIP-HFs’ hydrophobic environment. A medium exhibiting an intermediate polarity such as acetonitrile was also tested, resulting in recoveries lower than with methanol. In all cases, no analyte could be detected when the unmodified blank fibers (without MIP and NIP) were used as sorbent. In order to apply the prepared MIP-HFs in a relevant media, the recognition of the fibers previously conditioned in toluene, was tested in aqueous samples. This extraction mechanism would involve the liquid-liquid microextraction of the FQs to the toluene film impregnated in the walls of the fibers, followed by the subsequent recognition by the specific binding sites within the MIP. Although this approach showed a satisfactory recognition of triazines in water [23], in the current case the first barrier of toluene prevented the diffusion of the FQs to the MIP, according to their low octanol-water partition coefficient (log Pow ) [26]. Therefore, a different strategy was necessary to achieve specific recognition in aqueous medium. Fig. 4 shows the recovered CIP when including and increasing the percentage of water in methanol in the loading medium. Recoveries of MIP tend to decrease when increasing the water content into the loading solution. MIP-HFs exhibited a marked recovery drop with water percentages of 30% and higher. A water percentage of 20% was chosen as a compromise to keep an appropriate selective recognition without diluting excessively the aqueous sample.

As in any other extraction technique, parameters affecting the extraction efficiency include the extraction time and the energy provided to the system to promote the diffusion and partition of the analytes. On the one hand, the extraction time is of particular importance for equilibrium or non-exhaustive extraction techniques depending on the partition of the analytes and the donor/acceptor phases. On the other hand, the diffusion of the analytes is generally promoted by agitation and, it can be enhanced by heating or by tuning the composition of the phases (e.g. via a carrier-mediated extraction, modifying the polarity, the ionic strength of the media, etc). In the current case, a three level factorial design 32 involving 9 experiments was used to optimize the extraction time and the stirring rate. Spiked aqueous samples (0.8 mL) were diluted with methanol to a final volume of 4 mL so that the final dilution factor of the sample was 5. The spiking level of CIP was set at 0.5 mg L−1 for optimization. Fig. 5A includes the Pareto chart, which indicates that the response was mainly dependent on the speed of agitation, and to a lesser extent, also on the extraction time. The diffusion of the selected antibiotic is favored with a greater agitation, determining the amount of analyte that is able to bind the specific binding sites. The impact of the extraction time on the extraction efficiency indicates that the process operates out of equilibrium conditions. Fig. 5B shows the response surface for CIP obtained by the model extraction time and stirring rate. Based on these results, it was decided to perform the microextraction with a stirring speed of 750 rpm. As a compromise between extraction performance and duration of the process, the extraction time was set at 30 min.

3.4. Analytical performance and application to real samples The applicability of the MIP fibers to the analysis of FQs in real samples was studied and some analytical characteristics were evaluated. The prepared MIP-HFs were applied to the analysis of tap water samples spiked with CIP within the range 10–500 ␮g L−1 by HPLC-UV. The recoveries showed a linear tendency (r2 = 0.993) up to a loading concentration level of 100 ␮g L-1 , which can be considered the experimental loading capacity per MIP-HF. Fig. 6 shows clear differences between the polymer-HF capacity curves obtained for MIP and NIP, which confirms a different adsorption mechanism and therefore the presence of specific recognition sites in the MIP.

F. Barahona et al. / J. Chromatogr. A 1587 (2019) 42–49

Fig. 5. Pareto chart for ciprofloxacin showing the influence of each factor on the response (A) and response surface estimated from the factorial design by plotting extraction time versus stirring rate for ciprofloxacin (B).

Fig. 6. Amount of ciprofloxacin recovered by MIP and NIP-HFs when loading increasing concentrations of the analyte.

In order to lower the limits of detection (LOD), the analysis of spiked and real aqueous samples were carried out by means of HPLC-MS/MS as it is described in the methods section. Related target compounds, danofloxacin (DAN), norfloxacin (NOR), enrofloxacin (ENRO) and ciprofloxacin (CIP) were added to real liquid samples to test the applicability of the developed materials.

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Lake water and groundwater samples, polar liquid media where the presence of analytes can be expected, were spiked with the analytes. By visual inspection, the lake samples seemed to contain a greater amount of solid matter in suspension than the samples of groundwater. Similarly, due to the widespread use of these antibiotics, whether in animals or humans, their presence in biological samples can be also expected. Thus, a urine sample donated by a healthy volunteer was spiked with the four FQs as well. After oral ingestion, FQs are largely excreted by humans unaltered in urine high above mg L−1 concentration level in the 8 h after dose [29]. In the present work, original urine samples were diluted 10 times in pure water before mixing 1:5 with methanol. No further pre-treatment was applied to any replicate, and the developed MIPHFM methodology was directly applied for the analysis of urine samples. Calibrations by matrix matched standards were used to avoid matrix effects observed in HPLC-MS/MS analyses for each type of sample, generating calibration curves with correlation coefficients (R) that were: between 0.978 and 0.989 for surface water; between 0.979 and 0.990 for groundwater; and between 0.989 and 0,999 for urine samples. Fig. 7A shows the chromatogram by HPLC-MS/MS after the microextraction of the selected FQs in spiked urine samples. A detail of the magnification of the baseline signal is included in Fig. 7B. The chromatogram, obtained by monitoring the respective quantifying ions, shows the correct separation of all the selected antibiotics NOR, CIP, DAN and ENRO in an MIP-HFM extract of a spiked urine sample. A smooth flat baseline indicates that the proposed methodology generates clean extracts from water-based samples, allowing the determination of FQs in real samples of environmental and biological interest. The LODs for each compound and matrix were calculated as three times the signal to noise ratio in clean extracts. The LODs ranged between 2.0 and 10.0 ␮g L−1 in urine (for DAN and ENRO, respectively); 0.1 and 0.9 ␮g L−1 in surface waters (CIP and NOR, respectively); 0.6 and 2.4 ␮g L−1 in groundwater (DAN and CIP, respectively). Experimentally, it was possible to unequivocally quantify aqueous sample extracts spiked at 1 ␮g L−1 . Thanks to the clean-up capability of the polymer and to the greater sensitivity of the LC–MS/MS, the LODs improved those previously obtained for the analysis of FQs in soils [28] or infant foods [30] by means of HPLC-UV in a factor that ranged between 10 and 200, approximately. A detailed compilation of the analytical figures of merit obtained in the analyses of FQs in different types of samples is reviewed somewhere else [29]. The repeatability was studied in terms of the relative standard deviation (RSD) of the recoveries obtained from real samples spiked with the analytes at two different concentration levels, as shown in Table 2. In general, RSDs ranged between 2.1 and 19.1%, for all the analytes. Only in the case of groundwater at a lower spiking level, RSDs were above 20% for NOR and ENRO, fact that can be attributed to a likely instrumental deviation. These data indicate that both the preparation of the MIP composites and the developed MIP-HF microextraction procedure

Fig. 7. LC–MS/MS chromatograms obtained after the injection of an extract of spiked urine sample at 200 ␮g L−1 A) and a magnified detail of the baseline B).

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Table 2 Recoveries obtained from spiked aqueous samples at two different concentration levels. Relative standard deviations (RSD) were calculated for N = 3 (different individual fibers). Surface watera spiking level 100 ␮g L−1

NOR CIP DAN ENRO a b

Groundwatera spiking level 500 ␮g L−1

Urine sampleb

spiking level 100 ␮g L−1

spiking level 500 ␮g L−1

spiking level 200 ␮g L−1

spiking level 1000 ␮g L−1

R (%)

RSD

R (%)

RSD

R (%)

RSD

R (%)

RSD

R (%)

RSD

R (%)

RSD

13.9 17.6 16.6 17.0

2.1 13.4 15.4 11.3

18.0 18.0 26.0 15.7

4.3 5.2 2.5 2.6

10.1 16.1 12.4 10.4

24.8 18.4 17.8 22.3

16.7 19.1 16.2 15.1

6.4 14.8 8.8 8.6

23.5 24.5 12.5 16.5

19.1 16.3 20 18.1

18.4 15.6 9.4 19.5

6.0 10.2 13.8 5.1

Water samples were mixed with methanol 1:5 (v/v) before MIP-HFM (final dilution factor for water samples is 5). Urine samples was diluted 10 times in water, mixed with methanol 1:5 (v/v) before MIP-HFM (final dilution factor for urine samples is 50).

show acceptable repeatability. No apparent differences could be attributed among replicates from different type of sample. The analytical performance of the method depends to a large extent on the performance of the prepared fibers. The described polymerization process is homemade (lab-made) and adaptable to most laboratories. However, it can be easily automated to reduce the variability in fiber production, thus improving its analytical performance. The prepared MIP-HFs exhibited unaltered molecular recognition capability for approximately 15 microextraction cycles in real samples during the time of the experiments (c.a. three months). 4. Conclusions In this work, a new SPME fiber based on molecular imprinting technology has been proposed aiming to determine FQs in aqueous samples. First, a procedure to prepare a MIP immobilized in the pores of polypropylene hollow fibers has been developed and applied to the synthesis of MIP-HF composites. The polymerization process showed acceptable repeatability, both intra and inter-batch. Secondly, a microextraction methodology has been optimized using the resulting MIP-HFs to analyze FQs in real samples of environmental and biological interest. The MIP fibers exhibited selective recognition properties towards the selected hydrophilic antibiotics in toluene and in polar protic systems such as methanol and methanol/water solutions, up to 20% water content. This behavior is a remarkable feature, since it allows the application of the developed FQs-selective fibers to real waterbased samples. In addition, the methodology presented here also combines the advantages of miniaturized extraction techniques, such as a lower acceptor phase volume and a reduction in the use of organic solvents. Thus, working under optimal conditions it was possible to successfully apply the methodology to analyze real samples of surface water, groundwater and urine spiked with the analytes of interest. Acknowledgements Authors wish to thank the Spanish Ministry of Economy, Industry and Competitiveness for financial support, project (RTA2014-00012C03). Appendix A. Supplementary data Supplementary material related to this article can be found, in the online version, at doi:https://doi.org/10.1016/j.chroma.2018. 12.015. References [1] C.L. Arthur, J. Pawliszyn, Solid phase microextraction with thermal desorption using fused silica optical fibers, Anal. Chem. 62 (1990) 2145–2148, http://dx. doi.org/10.1021/ac00218a019.

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