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mass spectrometry
Microchemical Journal 110 (2013) 643–648
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Determination of parabens in waters by magnetically confined hydrophobic nanoparticle microextraction coupled to gas chromatography/mass spectrometry M.C. Alcudia-León, R. Lucena, S. Cárdenas, M. Valcárcel ⁎ Department of Analytical Chemistry, Institute of Fine Chemistry and Nanochemistry, Marie Curie Building, Campus de Rabanales, University of Córdoba, 14071 Córdoba, Spain
a r t i c l e
i n f o
Article history: Received 4 June 2013 Received in revised form 16 July 2013 Accepted 17 July 2013 Available online 24 July 2013 Keywords: Magnetically confined nanoparticles Octadecyl silica Parabens Sea water Swimming pool water
a b s t r a c t In this article, magnetically confined hydrophobic nanoparticle microextraction is applied for the analysis of parabens in water samples by gas chromatography (GC)–mass spectrometry (MS). The hydrophobic magnetic nanoparticles (MNPs) are confined in a device by means of a mini-magnet which also allows the stirring of the unit. A thin layer of MNPs, which presents an optimal surface to volume ratio, is the responsible of the analytes extraction. Although the superficial area of the layer is lower than the potential one obtained with a perfect dispersion of the MNPs, the latter approach is hard to be performed with highly hydrophobic MNPs and therefore the proposed configuration is more useful from a practical point of view. This fact, together with the inherent stirring of the unit, enhances the kinetic extraction and therefore the sensitivity of the procedure compared to conventional dispersion conditions. The most influential extraction parameters were evaluated, including the pH and ionic strength of the sample, the stirring rate, the extraction time, the amounts of MNPs, the volume of sample and the elution conditions. Under the most favorable extraction parameters, the method showed good linearity, repeatability (relative standard deviation below 7.1%, n = 7) and sensitivity in the ng per liter range. The proposed method was demonstrated to be a simple, fast and efficient method for the analysis of parabens in sea and swimming pool water samples. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Parabens are a group of alkyl esters of the p-hydroxybenzoic acid that are widely used as preservatives in pharmaceutical and personal care products (e.g.; solar lotions, deodorants, hair gels, shampoos, creams and toothpastes) due to their broad anti-microbial spectrum and effectiveness. Their anti-microbial activity increases with the length of the alkyl side chain from methyl to butyl. Parabens have multiple biological actions, but it is generally described that their inhibitory effects on membrane transport and mitochondrial function processes are keys for their actions [1]. Since their potential contribution to the incidence of breast cancer has been highlighted [2–6], the use of these preservatives in cosmetics has been discussed worldwide. As personal care products, parabens are continuously released into the environment through urban wastewater. Although they are relatively polar compounds that can be effectively removed during conventional sewage treatment [7–11], they have already been detected in surface waters at the ng/L level [9,10,12–14]. Moreover as parabens are usual components in sunscreen formulations, they can be present in sea water and swimming pool water. On the one hand, their occurrence in sea water is a problem of environmental concern ⁎ Corresponding author. Tel./fax: +34 957 218 616. E-mail address: [email protected] (M. Valcárcel). 0026-265X/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.microc.2013.07.011
since some parabens (especially butylparaben) can cause complete coral bleaching even at very low concentrations [15]. On the other hand, parabens can be found in swimming pool water as parent compounds and chlorinated by-products [16]. Thus, simple and sensitive analytical methods are required in order to understand the fate and distribution of this kind of emerging contaminants in the environment. The determination of parabens in water samples is usually accomplished by gas chromatography (GC) or liquid chromatography (LC). Due to the low concentration of the analytes in real samples, a previous preconcentration step is required. Solid phase extraction (SPE) is the common alternative of choice [9,10,17–22] since it allows for processing high sample volumes in a simple procedure. However, solid phase microextraction (SPME) has been firmly established as a valuable alternative to SPE methods [7,8] in the last years thanks to its low sample requirement to accomplish similar enrichment factors. Moreover, SPME requires a lower amount of organic solvents to be developed and it is easily coupled, even integrated, with commercial analytical instrumentation. In addition to SPE and SPME approaches, new microextraction techniques such as single-drop-microextraction (SDME) [23], ultrasound-assisted emulsification–microextraction [11] or membraneassisted liquid–liquid extraction [24] have been proposed to extract parabens from water samples. Recently a new extraction approach, called magnetically confined hydrophobic nanoparticle microextraction [25], has been proposed to
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take advantage of the special properties of octadecyl functionalized magnetic nanoparticles. This technique limits the aggregation tendency of these hydrophobic magnetic nanoparticles (MNPs) which can be considered their main drawback. The hydrophobic NPs are deposited in a special extraction device by means of a mini-magnet, which also allows the sample stirring in order to enhance the transference of the target analytes from the bulk sample to the extracting phase. The technique was evaluated for the determination of bisphenol A, 4-tertbutylphenol, 4-cumylphenol, 4-tert-octylphenol, 4-nonylphenol and 4octylphenol in water by LC. In this article, a new method for the extraction and determination of parabens in water samples is presented. The analytical procedure is based on a derivatization of the analytes with acetic anhydride, being the derivatized compounds finally isolated/preconcentrated by magnetically confined hydrophobic nanoparticle microextraction. The proposed method has been successfully applied to the determination of parabens in sea water and swimming pool water. 2. Experimental 2.1. Reagents, materials and samples All reagents were of analytical grade or better. Sigma-Aldrich (Madrid, Spain) provided the analytes: methyl 4-hydroxybenzoate (methyl-paraben), ethyl 4-hydroxybenzoate (ethyl-paraben), propyl 4-hydroxybenzoate (propyl-paraben) and butyl 4-hydroxybenzoate (butyl-paraben). Stock standard solutions of each analyte were prepared in methanol (Scharlab, Barcelona, Spain) at a concentration of 1 g/L and stored in the dark at 4 °C. Working solutions were prepared by a rigorous dilution of stock solutions with methanol or Milli-Q ultrapure water (Millipore Corp., Madrid, Spain) as required. Acetic anhydride and potassium carbonate, which were used for the analyte derivatization, were also purchased from Sigma-Aldrich. All the reagents required for the synthesis of the magnetic nanoparticles were also obtained from Sigma-Aldrich. Ferric chloride (FeCl3 · 6H2O), ferrous chloride (FeCl2 · 4H2O) and ammonia were used for the synthesis of the magnetic core (Fe3O4). Tetraethyl orthosilicate (TEOS) and ethanol were employed for covering the magnetic core with a protective silica-based coating. Finally, octadecyltriethoxysilane and toluene were used to introduce hydrophobic groups on the nanoparticle surface. Water samples, sea water (from the south coast of the Spain, collected near the shore) and swimming pool water, were collected in amberglass bottles without headspace. The samples were stored in the dark at 4 °C until their analysis. 2.2. Apparatus Gas chromatographic/mass spectrometric analyses were carried out on an HP6890 gas chromatograph equipped with an HP5973 (Agilent) mass spectrometric detector based on a quadrupole analyzer and an electron multiplier detector. System control and data acquisition was achieved with an HP1701CA MS ChemStation software (Agilent Technologies, Palo Alto, CA). Chromatographic separations were performed on a fused silica capillary column (30 m × 0.25 mm i.d.) coated with 5% diphenylsiloxane and 95% dimethylsiloxane (film thickness 0.25 μm) (Supelco, Madrid, Spain). The column temperature program was as follows: 1 min at 60 °C, raised up to 150 °C at 25 °C min−1, then immediately ramped at 3 °C min−1 up to 170 °C and raised up to 280 °C at 25 °C min−1 and kept finally at this temperature for 2 min. A column split ratio of 1:10 was selected for injection. Helium (6.0 grade purity, Air Liquide, Seville, Spain), at a flow rate of 1 mL/min, regulated by a digital pressure and flow controller, was used as carrier gas. Electron impact ionization (70 eV) was used for analyte fragmentation. The quadrupole mass spectrometer detector was operated in selected ion monitoring mode, recording 121 and 138 m/z ions. The
MS source and quadrupole temperatures were maintained at 230 and 150 °C, respectively. The peak areas were used for quantification of individual analytes. 2.3. Synthesis and characterization of magnetic nanoparticles MNPs are prepared as described in our previous work [25] following three main steps. First, the magnetic core (Fe304) was obtained by coprecipitation. For this aim, FeCl3·6H2O (24 g) and FeCl2·4H2O (9.8 g) were dissolved in 100 mL of water under nitrogen atmosphere, vigorously stirred and maintained at 80 °C in a water bath for 30 min. Then, 50 mL of ammonia (25 wt.%) were added dropwise producing a black precipitate of Fe3O4. The MNPs were separated with an external magnet, washed with water to remove the unreacted chemicals, and finally dried. In a second step, the MNPs were covered with a silica coating in order to protect the magnetite core. The Fe3O4@SiO2 nanoparticles were obtained by dispersing Fe3O4 nanoparticles in ethanol/water (50 mL/4 mL) solution in the presence of TEOS (2 mL) under a nitrogen atmosphere. The dispersion was stirred overnight and the protected magnetic nanoparticles were recovered with an external magnet, thoroughly washed with water and dried. Finally, the Fe3O4@SiO2 nanoparticles were dispersed in 50 mL of anhydrous toluene containing 1% (v/v) of octadecyltriethoxysilane. The mixture was sonicated for 5 min and refluxed for 12 h. The obtained nanoparticles (Fe3O4@SiO2@C18) were washed several times with ethanol and dried, yielding a fine powder of Fe3O4@SiO2@C18. The synthesized nanoparticles were characterized by FT-IR spectroscopy and microscopy. The particle size and structure of MNPs were observed by using a transmission electron microscopy (TEM). They show a spherical morphology and an average particle size of 10 nm. The IR spectrum obtained under the attenuated total reflection sampling mode shows a characteristic band of Fe3O4 around 600 cm−1 which corresponds to the Fe–O bonds and a strong absorbing region at 1200–1000 cm−1 characteristic of the Si–O–H and Si–O–Si bonds. Moreover, the characteristics C\H stretching vibrating bands of octadecyl groups of the Fe3O4@SiO2@C18 MNPs can be observed at 2920 and 2850 cm−1. 2.4. Magnetically confined MNPs unit design The extraction unit is described elsewhere [25] and it consists of four commercial elements, namely: (a) a cube-shaped magnet (5 mm in length and 10.8 N of maximum adhesive force) purchased from Supermagnete (Gottmadingen, Germany), (b) a PTFE septum (c) a PTFE top-cap commercially available for the SPE cartridges, and (d) an
Fig. 1. Schematic view of the extraction device.
M.C. Alcudia-León et al. / Microchemical Journal 110 (2013) 643–648 Table 1 List of the variables involved in the proposed extraction technique. Variable Stirring rate (rpm) Stirring time (min) MNPs amount (mg) Sample volume (mL)
Initial value 10 5 25
Interval studied
Optimal value
0–1600 2–60 2–30 15–100
1400 20 10 30 Ethyl acetate
Eluent
Methanol
Methanol, acetonitrile, acetone, toluene, ethyl acetate
Eluent volume (μL) Elution stirring rate (rpm) Elution time (min)
100 300 2
50–150 0–500 1–5
100 100 2
external part, which is the upper section of a commercial 3 mL SPE cartridge (1 cm internal diameter, 1 cm height). In the assembly process, the magnet is introduced in the PTFE top cap which is subsequently sealed with the septum. In this way the magnet, which plays a double function since it allows the stirring of the unit and the confinement of the magnetic nanoparticles, is protected from the sample matrix. Finally, the upper section of the solid phase extraction cartridge is displaced through the PTFE top cap. Due to the difference in height (10 mm), a small chamber is then obtained which will contain the Fe3O4@SiO2@C18 nanoparticles during the extraction. A schematic view of the device is depicted in Fig. 1. 2.5. Extraction procedure In order to make easier the extraction and gas chromatographic analysis of the target analytes, they should be previously derivatized. For this purpose, 30 mL of the aqueous standard or the sample containing the selected parabens are placed in 50 mL beaker and 0.6 mL of 5 M K2CO3 solution and 0.6 mL of acetic anhydride (derivatization reagent) are added. The solution is stirred for 2 min in a magnetic stirrer (500 rpm). Meanwhile, the extraction unit containing 10 mg of the Fe3O4@SiO2@C18 MNPs is subsequently conditioned with methanol (1 min) and Milli-Q water (1 min). Then, the unit is introduced into the aqueous standard or sample containing the derivatized analytes. The beaker is placed on a magnetic stirrer and the unit is stirred at 1400 rpm for 20 min. After the extraction, the unit is withdrawn from the beaker and directly placed on the magnetic stirrer for analyte
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elution. The parabens were desorbed from the Fe3O4@SiO2@C18 MNPs with 100 μL of ethyl acetate under continuous magnetic stirring of the unit (100 rpm) for 2 min. The organic extract is recovered and placed into a 200 μL-insert of a 1.5 mL vial for further GC analysis. After the extraction, the unit just required a soft cleaning with methanol and water and it can be reused. 3. Results and discussion 3.1. Optimization of the extraction conditions A general solid phase extraction procedure may be affected by several variables such as the pH, ionic strength, amount of sorbent, sample volume, stirring time, extraction time and elution conditions. The first two variables do not present a relevant effect on the extraction of parabens by the magnetically confined hydrophobic nanoparticles microextraction due to the derivatization process. On the one hand, derivatized analytes do not present any ionizable groups and therefore the sample pH is not critical in their extraction. On the other hand, an excess of electrolytes is added during the derivatization process which makes negligible the effect of the intrinsic ionic strength of the sample. A univariate method was employed to optimize the proposed procedure. The studied variables, including their initial values, are shown in Table 1. The effect of the stirring rate on the extraction efficiency was studied within the interval 0–1600 rpm. The results, which are summarized in Fig. 2, show an increase of the signal with the stirring rate up to 1400 rpm. This fact can be ascribed to the enhancement of the analyte transfer from the bulk solution to the extracting phase. However, a signal decrease was observed for higher rates probably due to the sorbent loss from the extraction device. Considering the results 1400 rpm was selected as the optimum value. Extraction time is a key parameter affecting the method sensitivity in some extraction techniques. This variable was studied in the interval from 2 to 60 min, the result being shown in Fig. 3. A rapid increase in the analytical signals was obtained for the four parabens between 2 and 20 min. After this time, the extraction efficiency did not change significantly for most of the analytes and therefore 20 min was chosen as the optimum values. The obtained MNPs show high adsorption towards the derivatized parabens due to their high surface area and the presence of hydrophobic C18 chains on their surface. The influence of the MNP amount has an
Fig. 2. Effect of the stirring rate on the extraction of the target analytes.
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Fig. 3. Effect of the extraction time on the extraction of the target analytes.
expected influence on the results. The analytical signal increases with the MNP amount up to 10 mg due to the inherent increase of sorption capacity. At higher amount of MNPs a decrease in the analytical signal is observed. This fact can be ascribed to two complementary reasons. On the one hand, the use of higher amounts of MNPs induces the formation of a thicker MNP layer which is not practical for extraction purposes since only the superficial layer can interact easily with the analytes. On the other hand, the detachment of the MNPs from the extraction unit is more probable when high amounts of the sorbent are employed. According to the results, 10 mg was selected as the optimum value. The sample volume is a critical parameter in the extraction procedure since it defines the maximum enrichment factor that can be reached in the procedure. The effect of donor phase volume (standard or sample) was examined in the interval 15–100 mL at 10 μg L−1 level. As can be observed in Fig. 4, the analyte extraction was maximum for 30 mL at both concentration levels, remaining almost constant for higher volumes. Therefore, 30 mL was selected as the sample volume for further optimization of the method.
Finally, different organic solvents (methanol, acetonitrile, acetone, toluene, ethyl acetate) were evaluated as eluents, the best result being obtained from 100 μL of ethyl acetate (see Fig. 5). Concerning the elution conditions, satisfactory results were obtained when elution was performed at 100 rpm for 2 min. 3.2. Analytical figures of merit After the optimization process, the proposed procedure was evaluated in terms of linearity, sensitivity, precision and accuracy. The first three analytical features are summarized in Table 2. The data were obtained after a calibration of the proposed method by analyzing a series of standard solutions containing the four parabens under the optimized operational condition. Each standard was extracted under the optimum conditions. The linear range was 0.5–500 μg L−1 for methylparaben, while the linearity value was in the interval from 0.1 to 500 μg L−1 for ethyl, propyl and butyl-paraben. The correlation coefficients (R2) were in the range 0.996–0.999 for all analytes. The limits of
Fig. 4. Effect of the sample volume on the extraction of the target analytes.
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Fig. 5. Effect of the type of eluent on the analytical signal.
detection, calculated on the basis of signal to noise ratio of 3 (S/N = 3), varied between 23.2 ng L−1 (for butyl-paraben) and 86.1 ng L−1 (for methyl-paraben). The precision of the method, expressed as relative standard deviation (RSD) was evaluated by analyzing 7 individual aqueous standards at a concentration of 0.5 μg L−1. Once optimized and analytically characterized, the proposed method was applied to the determination of the target analytes in 2 swimming pool water and 7 sea water samples, where these analytes are expected to be found due to the widespread use of solar lotions. Samples were analyzed in triplicate and they were directly extracted using the developed method without any pretreatment. Butyl-paraben was detected in six samples (1 swimming pool and 5 sea water) and propyl-paraben was detected in only one sea water sample. Although the analytes were detected in some samples their concentrations were below the quantification limit and therefore no quantitative results can be provided. In order to evaluate the suitability of the proposed method for the determination of the analyte in the sample, a recovery Table 2 Analytical figures of merit of the proposed method. Analyte
Linear range (μg L−1)
R2
LODa (ng L−1)
RSDb (%)
Methyl-paraben Ethyl-paraben Propyl-paraben Butyl-paraben
0.5–500 0.1–500 0.1–500 0.1–500
0.998 0.996 0.999 0.999
86.1 34.6 23.8 23.2
7.1 6.9 5.1 6.5
a b
LOD, limit of detection. RSD, relative standard deviation (n = 7).
study at two concentration levels (0.5 and 5 μg L−1) was accomplished. The mean recoveries for all analytes were in the range of 96–106%, which indicated that the method was reliable and can be used for the determination of parabens in water samples. The results are summarized in Table 3. 4. Conclusions In the present work, the potential of microextraction with magnetically confined hydrophobic nanoparticles has been evaluated as a sample preparation technique for the extraction of parabens from swimming pool and sea water samples. All the variables involved in the extraction procedure were studied and optimized. Furthermore, the methodology was successfully evaluated in terms of linearity, sensitivity (limits of detection and quantification), precision and accuracy (recovery studies). The developed extraction technique allows the efficient isolation and preconcentration of the target analytes and its combination with GC/MS has been demonstrated to be viable, easy to use, rapid and economical. Some improvements would still be accomplished by using, for example, other surface modified MNPs with other types of extraction group to assist in the extraction of the target analytes. Nevertheless, the findings in this work indicate that this procedure can be applied for enrichment of parabens from complicated matrices such as sea water. Table 4 presents a critical comparison of the proposed method with other counterparts [8,12,23,24,26–31]. In terms of sensitivity, the new
Table 3 Relative recoveries obtained for the application of of parabens in seawater and swimming pool water samples. Sea I
Sea II −1
−1
Swimming pool I −1
−1
Analyte
0.5 μg L
5 μg L
0.5 μg L
5 μg L
Methyl-paraben Ethyl-paraben Propyl-paraben Butyl-paraben
103 ± 106 ± 103 ± 101 ±
104 104 102 102
103 ± 7 104 ± 7 103 ± 5 104 ± 7
99 ± 101 ± 100 ± 99 ±
7 7 5 7
±7 ±7 ±5 ±7
7 7 5 6
−1
Swimming pool II −1
0.5 μg L
5 μg L
0.5 μg L−1
5 μg L−1
101 ± 7 100 ± 7 101 ± 5 100 ± 7
97 99 98 99
96 ± 98 ± 98 ± 101 ±
103 101 102 99
±7 ±7 ±5 ±6
7 7 5 7
±7 ±7 ±5 ±6
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Table 4 Comparison of the proposed method with other published alternatives for the determination of parabens in aqueous matrices. Extraction techniquea
Samples
Instrumental LODc techniqueb (ng/L)
RSDd (%)
Reference
SPME MEPS SDME Membrane assisted LLE DLLME
River water Wastewater River water Wastewater
GC–MS-MS LVI-GC–MS GC–MS LVI-GC-ion trap MS GC-FID
1–25 20–590 1–15 0.1–1.4
b11.9 b7.1% b13 b6.8
[8] [12] [23] [24]
29–102
b6.86 [26]
GC-PID HPLC-UV HPLC-UV HPLC-UV TD-GC–MS GC–MS
50–300 50–200 21–46 300–400 0.03–0.3 23.2–86.1
Solvent assisted DSPE Emulsification LPME DLLME Magnetic SPE SBSE Magnetically confined hydrophobic nanoparticles microextraction
Food and water River water Water Tap water Wastewater Wastewater Swimming pool and seawater samples
b8 b5.5 b9.3 b2.4 b13 b7.1
[27] [28] [29] [30] [31] This work
a Acronyms: SPME, solid phase microextraction; MEPS, microextraction by packed sorbent; SDME, single drop microextraction; LLE, liquid–liquid extraction; DLLME, dispersive liquid–liquid extraction; DSPE, dispersive solid phase extraction, LPME, liquid phase microextraction; SBSE, stir bar sorptive extraction. b Acronyms: GC, gas chromatography; MS, mass spectrometry; LVI, large volume injection; FID, flame ionization detector; PID, photo-ionization detector, HPLC, high performance liquid chromatography; UV, ultraviolet detection, TD, thermal desorption. c LOD, limit of detection. d RSD, relative standard deviation.
approach provides similar LODs than the obtained with dispersive liquid–liquid microextraction (DLLME) [26,29] and better results than those obtained with microextraction by packed sorbents (MEPS) [12], solvent assisted dispersive SPE [27], emulsification liquid phase microextraction [28] and magnetic SPE [30]. However, some of the latter approaches [12,30] have been successfully applied to the analysis of wastewater samples. Moreover, the rest of the methodologies [8,23,24,31] are more sensitive due to different reasons. The use of SPME [8] or SDME [23] involves a sensitivity enhancement since all the extracts (the fiber or the single drop) can be injected without a previous elution. On the other hand, the use of large volume injection (LVI) [24] or thermal desorption [31] allows the introduction of higher amounts of analytes in the instrument, improving also the sensitivity of the method. Concerning the precision, our method provides the fifth best value. Acknowledgments Financial support from the Spanish Ministry of Science and Innovation (grant CTQ2011-23790) and P09-FQM-4801 from the Junta de Andalucía are gratefully acknowledged. References [1] P.D. Darbre, A. Aljarrah, W.R. Miller, N.G. Coldham, M.J. Sauer, G.S. Pope, Concentrations of parabens in human breast tumours, J. Appl. Toxicol. 24 (2004) 5–13. [2] P.D. Darbre, Underarm cosmetics and breast cancer, J. Appl. Toxicol. 23 (2003) 89–95. [3] P.D. Darbre, P.W. Harvey, Paraben esters: review of recent studies of endocrine toxicity, absorption, esterase and human exposure, and discussion of potential human health risks, J. Appl. Toxicol. 28 (2008) 561–578. [4] E.J. Routledge, J. Parker, J. Odum, J. Ashby, J.P. Sumpter, Some alkyl hydroxyl benzoate preservatives (parabens) are estrogenic, Toxicol. Appl. Pharmacol. 153 (1998) 12–19. [5] R. Golden, J. Gandy, G. Vollmer, A review of the endocrine activity of parabens and implications for potential risks to human health, Crit. Rev. Toxicol. 35 (2005) 435–458. [6] J.M. Brausch, G.M. Rand, A review of personal care products in the aquatic environment: environmental concentrations and toxicity, Chemosphere 82 (2011) 1518–1532.
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Determination of parabens in waters by magnetically confined hydrophobic nanoparticle microextraction coupled to gas chromatography/mass spectrometry M.C. Alcudia-León, R. Lucena, S. Cárdenas, M. Valcárcel ⁎ Department of Analytical Chemistry, Institute of Fine Chemistry and Nanochemistry, Marie Curie Building, Campus de Rabanales, University of Córdoba, 14071 Córdoba, Spain
a r t i c l e
i n f o
Article history: Received 4 June 2013 Received in revised form 16 July 2013 Accepted 17 July 2013 Available online 24 July 2013 Keywords: Magnetically confined nanoparticles Octadecyl silica Parabens Sea water Swimming pool water
a b s t r a c t In this article, magnetically confined hydrophobic nanoparticle microextraction is applied for the analysis of parabens in water samples by gas chromatography (GC)–mass spectrometry (MS). The hydrophobic magnetic nanoparticles (MNPs) are confined in a device by means of a mini-magnet which also allows the stirring of the unit. A thin layer of MNPs, which presents an optimal surface to volume ratio, is the responsible of the analytes extraction. Although the superficial area of the layer is lower than the potential one obtained with a perfect dispersion of the MNPs, the latter approach is hard to be performed with highly hydrophobic MNPs and therefore the proposed configuration is more useful from a practical point of view. This fact, together with the inherent stirring of the unit, enhances the kinetic extraction and therefore the sensitivity of the procedure compared to conventional dispersion conditions. The most influential extraction parameters were evaluated, including the pH and ionic strength of the sample, the stirring rate, the extraction time, the amounts of MNPs, the volume of sample and the elution conditions. Under the most favorable extraction parameters, the method showed good linearity, repeatability (relative standard deviation below 7.1%, n = 7) and sensitivity in the ng per liter range. The proposed method was demonstrated to be a simple, fast and efficient method for the analysis of parabens in sea and swimming pool water samples. © 2013 Elsevier B.V. All rights reserved.
1. Introduction Parabens are a group of alkyl esters of the p-hydroxybenzoic acid that are widely used as preservatives in pharmaceutical and personal care products (e.g.; solar lotions, deodorants, hair gels, shampoos, creams and toothpastes) due to their broad anti-microbial spectrum and effectiveness. Their anti-microbial activity increases with the length of the alkyl side chain from methyl to butyl. Parabens have multiple biological actions, but it is generally described that their inhibitory effects on membrane transport and mitochondrial function processes are keys for their actions [1]. Since their potential contribution to the incidence of breast cancer has been highlighted [2–6], the use of these preservatives in cosmetics has been discussed worldwide. As personal care products, parabens are continuously released into the environment through urban wastewater. Although they are relatively polar compounds that can be effectively removed during conventional sewage treatment [7–11], they have already been detected in surface waters at the ng/L level [9,10,12–14]. Moreover as parabens are usual components in sunscreen formulations, they can be present in sea water and swimming pool water. On the one hand, their occurrence in sea water is a problem of environmental concern ⁎ Corresponding author. Tel./fax: +34 957 218 616. E-mail address: [email protected] (M. Valcárcel). 0026-265X/$ – see front matter © 2013 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.microc.2013.07.011
since some parabens (especially butylparaben) can cause complete coral bleaching even at very low concentrations [15]. On the other hand, parabens can be found in swimming pool water as parent compounds and chlorinated by-products [16]. Thus, simple and sensitive analytical methods are required in order to understand the fate and distribution of this kind of emerging contaminants in the environment. The determination of parabens in water samples is usually accomplished by gas chromatography (GC) or liquid chromatography (LC). Due to the low concentration of the analytes in real samples, a previous preconcentration step is required. Solid phase extraction (SPE) is the common alternative of choice [9,10,17–22] since it allows for processing high sample volumes in a simple procedure. However, solid phase microextraction (SPME) has been firmly established as a valuable alternative to SPE methods [7,8] in the last years thanks to its low sample requirement to accomplish similar enrichment factors. Moreover, SPME requires a lower amount of organic solvents to be developed and it is easily coupled, even integrated, with commercial analytical instrumentation. In addition to SPE and SPME approaches, new microextraction techniques such as single-drop-microextraction (SDME) [23], ultrasound-assisted emulsification–microextraction [11] or membraneassisted liquid–liquid extraction [24] have been proposed to extract parabens from water samples. Recently a new extraction approach, called magnetically confined hydrophobic nanoparticle microextraction [25], has been proposed to
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take advantage of the special properties of octadecyl functionalized magnetic nanoparticles. This technique limits the aggregation tendency of these hydrophobic magnetic nanoparticles (MNPs) which can be considered their main drawback. The hydrophobic NPs are deposited in a special extraction device by means of a mini-magnet, which also allows the sample stirring in order to enhance the transference of the target analytes from the bulk sample to the extracting phase. The technique was evaluated for the determination of bisphenol A, 4-tertbutylphenol, 4-cumylphenol, 4-tert-octylphenol, 4-nonylphenol and 4octylphenol in water by LC. In this article, a new method for the extraction and determination of parabens in water samples is presented. The analytical procedure is based on a derivatization of the analytes with acetic anhydride, being the derivatized compounds finally isolated/preconcentrated by magnetically confined hydrophobic nanoparticle microextraction. The proposed method has been successfully applied to the determination of parabens in sea water and swimming pool water. 2. Experimental 2.1. Reagents, materials and samples All reagents were of analytical grade or better. Sigma-Aldrich (Madrid, Spain) provided the analytes: methyl 4-hydroxybenzoate (methyl-paraben), ethyl 4-hydroxybenzoate (ethyl-paraben), propyl 4-hydroxybenzoate (propyl-paraben) and butyl 4-hydroxybenzoate (butyl-paraben). Stock standard solutions of each analyte were prepared in methanol (Scharlab, Barcelona, Spain) at a concentration of 1 g/L and stored in the dark at 4 °C. Working solutions were prepared by a rigorous dilution of stock solutions with methanol or Milli-Q ultrapure water (Millipore Corp., Madrid, Spain) as required. Acetic anhydride and potassium carbonate, which were used for the analyte derivatization, were also purchased from Sigma-Aldrich. All the reagents required for the synthesis of the magnetic nanoparticles were also obtained from Sigma-Aldrich. Ferric chloride (FeCl3 · 6H2O), ferrous chloride (FeCl2 · 4H2O) and ammonia were used for the synthesis of the magnetic core (Fe3O4). Tetraethyl orthosilicate (TEOS) and ethanol were employed for covering the magnetic core with a protective silica-based coating. Finally, octadecyltriethoxysilane and toluene were used to introduce hydrophobic groups on the nanoparticle surface. Water samples, sea water (from the south coast of the Spain, collected near the shore) and swimming pool water, were collected in amberglass bottles without headspace. The samples were stored in the dark at 4 °C until their analysis. 2.2. Apparatus Gas chromatographic/mass spectrometric analyses were carried out on an HP6890 gas chromatograph equipped with an HP5973 (Agilent) mass spectrometric detector based on a quadrupole analyzer and an electron multiplier detector. System control and data acquisition was achieved with an HP1701CA MS ChemStation software (Agilent Technologies, Palo Alto, CA). Chromatographic separations were performed on a fused silica capillary column (30 m × 0.25 mm i.d.) coated with 5% diphenylsiloxane and 95% dimethylsiloxane (film thickness 0.25 μm) (Supelco, Madrid, Spain). The column temperature program was as follows: 1 min at 60 °C, raised up to 150 °C at 25 °C min−1, then immediately ramped at 3 °C min−1 up to 170 °C and raised up to 280 °C at 25 °C min−1 and kept finally at this temperature for 2 min. A column split ratio of 1:10 was selected for injection. Helium (6.0 grade purity, Air Liquide, Seville, Spain), at a flow rate of 1 mL/min, regulated by a digital pressure and flow controller, was used as carrier gas. Electron impact ionization (70 eV) was used for analyte fragmentation. The quadrupole mass spectrometer detector was operated in selected ion monitoring mode, recording 121 and 138 m/z ions. The
MS source and quadrupole temperatures were maintained at 230 and 150 °C, respectively. The peak areas were used for quantification of individual analytes. 2.3. Synthesis and characterization of magnetic nanoparticles MNPs are prepared as described in our previous work [25] following three main steps. First, the magnetic core (Fe304) was obtained by coprecipitation. For this aim, FeCl3·6H2O (24 g) and FeCl2·4H2O (9.8 g) were dissolved in 100 mL of water under nitrogen atmosphere, vigorously stirred and maintained at 80 °C in a water bath for 30 min. Then, 50 mL of ammonia (25 wt.%) were added dropwise producing a black precipitate of Fe3O4. The MNPs were separated with an external magnet, washed with water to remove the unreacted chemicals, and finally dried. In a second step, the MNPs were covered with a silica coating in order to protect the magnetite core. The Fe3O4@SiO2 nanoparticles were obtained by dispersing Fe3O4 nanoparticles in ethanol/water (50 mL/4 mL) solution in the presence of TEOS (2 mL) under a nitrogen atmosphere. The dispersion was stirred overnight and the protected magnetic nanoparticles were recovered with an external magnet, thoroughly washed with water and dried. Finally, the Fe3O4@SiO2 nanoparticles were dispersed in 50 mL of anhydrous toluene containing 1% (v/v) of octadecyltriethoxysilane. The mixture was sonicated for 5 min and refluxed for 12 h. The obtained nanoparticles (Fe3O4@SiO2@C18) were washed several times with ethanol and dried, yielding a fine powder of Fe3O4@SiO2@C18. The synthesized nanoparticles were characterized by FT-IR spectroscopy and microscopy. The particle size and structure of MNPs were observed by using a transmission electron microscopy (TEM). They show a spherical morphology and an average particle size of 10 nm. The IR spectrum obtained under the attenuated total reflection sampling mode shows a characteristic band of Fe3O4 around 600 cm−1 which corresponds to the Fe–O bonds and a strong absorbing region at 1200–1000 cm−1 characteristic of the Si–O–H and Si–O–Si bonds. Moreover, the characteristics C\H stretching vibrating bands of octadecyl groups of the Fe3O4@SiO2@C18 MNPs can be observed at 2920 and 2850 cm−1. 2.4. Magnetically confined MNPs unit design The extraction unit is described elsewhere [25] and it consists of four commercial elements, namely: (a) a cube-shaped magnet (5 mm in length and 10.8 N of maximum adhesive force) purchased from Supermagnete (Gottmadingen, Germany), (b) a PTFE septum (c) a PTFE top-cap commercially available for the SPE cartridges, and (d) an
Fig. 1. Schematic view of the extraction device.
M.C. Alcudia-León et al. / Microchemical Journal 110 (2013) 643–648 Table 1 List of the variables involved in the proposed extraction technique. Variable Stirring rate (rpm) Stirring time (min) MNPs amount (mg) Sample volume (mL)
Initial value 10 5 25
Interval studied
Optimal value
0–1600 2–60 2–30 15–100
1400 20 10 30 Ethyl acetate
Eluent
Methanol
Methanol, acetonitrile, acetone, toluene, ethyl acetate
Eluent volume (μL) Elution stirring rate (rpm) Elution time (min)
100 300 2
50–150 0–500 1–5
100 100 2
external part, which is the upper section of a commercial 3 mL SPE cartridge (1 cm internal diameter, 1 cm height). In the assembly process, the magnet is introduced in the PTFE top cap which is subsequently sealed with the septum. In this way the magnet, which plays a double function since it allows the stirring of the unit and the confinement of the magnetic nanoparticles, is protected from the sample matrix. Finally, the upper section of the solid phase extraction cartridge is displaced through the PTFE top cap. Due to the difference in height (10 mm), a small chamber is then obtained which will contain the Fe3O4@SiO2@C18 nanoparticles during the extraction. A schematic view of the device is depicted in Fig. 1. 2.5. Extraction procedure In order to make easier the extraction and gas chromatographic analysis of the target analytes, they should be previously derivatized. For this purpose, 30 mL of the aqueous standard or the sample containing the selected parabens are placed in 50 mL beaker and 0.6 mL of 5 M K2CO3 solution and 0.6 mL of acetic anhydride (derivatization reagent) are added. The solution is stirred for 2 min in a magnetic stirrer (500 rpm). Meanwhile, the extraction unit containing 10 mg of the Fe3O4@SiO2@C18 MNPs is subsequently conditioned with methanol (1 min) and Milli-Q water (1 min). Then, the unit is introduced into the aqueous standard or sample containing the derivatized analytes. The beaker is placed on a magnetic stirrer and the unit is stirred at 1400 rpm for 20 min. After the extraction, the unit is withdrawn from the beaker and directly placed on the magnetic stirrer for analyte
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elution. The parabens were desorbed from the Fe3O4@SiO2@C18 MNPs with 100 μL of ethyl acetate under continuous magnetic stirring of the unit (100 rpm) for 2 min. The organic extract is recovered and placed into a 200 μL-insert of a 1.5 mL vial for further GC analysis. After the extraction, the unit just required a soft cleaning with methanol and water and it can be reused. 3. Results and discussion 3.1. Optimization of the extraction conditions A general solid phase extraction procedure may be affected by several variables such as the pH, ionic strength, amount of sorbent, sample volume, stirring time, extraction time and elution conditions. The first two variables do not present a relevant effect on the extraction of parabens by the magnetically confined hydrophobic nanoparticles microextraction due to the derivatization process. On the one hand, derivatized analytes do not present any ionizable groups and therefore the sample pH is not critical in their extraction. On the other hand, an excess of electrolytes is added during the derivatization process which makes negligible the effect of the intrinsic ionic strength of the sample. A univariate method was employed to optimize the proposed procedure. The studied variables, including their initial values, are shown in Table 1. The effect of the stirring rate on the extraction efficiency was studied within the interval 0–1600 rpm. The results, which are summarized in Fig. 2, show an increase of the signal with the stirring rate up to 1400 rpm. This fact can be ascribed to the enhancement of the analyte transfer from the bulk solution to the extracting phase. However, a signal decrease was observed for higher rates probably due to the sorbent loss from the extraction device. Considering the results 1400 rpm was selected as the optimum value. Extraction time is a key parameter affecting the method sensitivity in some extraction techniques. This variable was studied in the interval from 2 to 60 min, the result being shown in Fig. 3. A rapid increase in the analytical signals was obtained for the four parabens between 2 and 20 min. After this time, the extraction efficiency did not change significantly for most of the analytes and therefore 20 min was chosen as the optimum values. The obtained MNPs show high adsorption towards the derivatized parabens due to their high surface area and the presence of hydrophobic C18 chains on their surface. The influence of the MNP amount has an
Fig. 2. Effect of the stirring rate on the extraction of the target analytes.
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Fig. 3. Effect of the extraction time on the extraction of the target analytes.
expected influence on the results. The analytical signal increases with the MNP amount up to 10 mg due to the inherent increase of sorption capacity. At higher amount of MNPs a decrease in the analytical signal is observed. This fact can be ascribed to two complementary reasons. On the one hand, the use of higher amounts of MNPs induces the formation of a thicker MNP layer which is not practical for extraction purposes since only the superficial layer can interact easily with the analytes. On the other hand, the detachment of the MNPs from the extraction unit is more probable when high amounts of the sorbent are employed. According to the results, 10 mg was selected as the optimum value. The sample volume is a critical parameter in the extraction procedure since it defines the maximum enrichment factor that can be reached in the procedure. The effect of donor phase volume (standard or sample) was examined in the interval 15–100 mL at 10 μg L−1 level. As can be observed in Fig. 4, the analyte extraction was maximum for 30 mL at both concentration levels, remaining almost constant for higher volumes. Therefore, 30 mL was selected as the sample volume for further optimization of the method.
Finally, different organic solvents (methanol, acetonitrile, acetone, toluene, ethyl acetate) were evaluated as eluents, the best result being obtained from 100 μL of ethyl acetate (see Fig. 5). Concerning the elution conditions, satisfactory results were obtained when elution was performed at 100 rpm for 2 min. 3.2. Analytical figures of merit After the optimization process, the proposed procedure was evaluated in terms of linearity, sensitivity, precision and accuracy. The first three analytical features are summarized in Table 2. The data were obtained after a calibration of the proposed method by analyzing a series of standard solutions containing the four parabens under the optimized operational condition. Each standard was extracted under the optimum conditions. The linear range was 0.5–500 μg L−1 for methylparaben, while the linearity value was in the interval from 0.1 to 500 μg L−1 for ethyl, propyl and butyl-paraben. The correlation coefficients (R2) were in the range 0.996–0.999 for all analytes. The limits of
Fig. 4. Effect of the sample volume on the extraction of the target analytes.
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Fig. 5. Effect of the type of eluent on the analytical signal.
detection, calculated on the basis of signal to noise ratio of 3 (S/N = 3), varied between 23.2 ng L−1 (for butyl-paraben) and 86.1 ng L−1 (for methyl-paraben). The precision of the method, expressed as relative standard deviation (RSD) was evaluated by analyzing 7 individual aqueous standards at a concentration of 0.5 μg L−1. Once optimized and analytically characterized, the proposed method was applied to the determination of the target analytes in 2 swimming pool water and 7 sea water samples, where these analytes are expected to be found due to the widespread use of solar lotions. Samples were analyzed in triplicate and they were directly extracted using the developed method without any pretreatment. Butyl-paraben was detected in six samples (1 swimming pool and 5 sea water) and propyl-paraben was detected in only one sea water sample. Although the analytes were detected in some samples their concentrations were below the quantification limit and therefore no quantitative results can be provided. In order to evaluate the suitability of the proposed method for the determination of the analyte in the sample, a recovery Table 2 Analytical figures of merit of the proposed method. Analyte
Linear range (μg L−1)
R2
LODa (ng L−1)
RSDb (%)
Methyl-paraben Ethyl-paraben Propyl-paraben Butyl-paraben
0.5–500 0.1–500 0.1–500 0.1–500
0.998 0.996 0.999 0.999
86.1 34.6 23.8 23.2
7.1 6.9 5.1 6.5
a b
LOD, limit of detection. RSD, relative standard deviation (n = 7).
study at two concentration levels (0.5 and 5 μg L−1) was accomplished. The mean recoveries for all analytes were in the range of 96–106%, which indicated that the method was reliable and can be used for the determination of parabens in water samples. The results are summarized in Table 3. 4. Conclusions In the present work, the potential of microextraction with magnetically confined hydrophobic nanoparticles has been evaluated as a sample preparation technique for the extraction of parabens from swimming pool and sea water samples. All the variables involved in the extraction procedure were studied and optimized. Furthermore, the methodology was successfully evaluated in terms of linearity, sensitivity (limits of detection and quantification), precision and accuracy (recovery studies). The developed extraction technique allows the efficient isolation and preconcentration of the target analytes and its combination with GC/MS has been demonstrated to be viable, easy to use, rapid and economical. Some improvements would still be accomplished by using, for example, other surface modified MNPs with other types of extraction group to assist in the extraction of the target analytes. Nevertheless, the findings in this work indicate that this procedure can be applied for enrichment of parabens from complicated matrices such as sea water. Table 4 presents a critical comparison of the proposed method with other counterparts [8,12,23,24,26–31]. In terms of sensitivity, the new
Table 3 Relative recoveries obtained for the application of of parabens in seawater and swimming pool water samples. Sea I
Sea II −1
−1
Swimming pool I −1
−1
Analyte
0.5 μg L
5 μg L
0.5 μg L
5 μg L
Methyl-paraben Ethyl-paraben Propyl-paraben Butyl-paraben
103 ± 106 ± 103 ± 101 ±
104 104 102 102
103 ± 7 104 ± 7 103 ± 5 104 ± 7
99 ± 101 ± 100 ± 99 ±
7 7 5 7
±7 ±7 ±5 ±7
7 7 5 6
−1
Swimming pool II −1
0.5 μg L
5 μg L
0.5 μg L−1
5 μg L−1
101 ± 7 100 ± 7 101 ± 5 100 ± 7
97 99 98 99
96 ± 98 ± 98 ± 101 ±
103 101 102 99
±7 ±7 ±5 ±6
7 7 5 7
±7 ±7 ±5 ±6
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Table 4 Comparison of the proposed method with other published alternatives for the determination of parabens in aqueous matrices. Extraction techniquea
Samples
Instrumental LODc techniqueb (ng/L)
RSDd (%)
Reference
SPME MEPS SDME Membrane assisted LLE DLLME
River water Wastewater River water Wastewater
GC–MS-MS LVI-GC–MS GC–MS LVI-GC-ion trap MS GC-FID
1–25 20–590 1–15 0.1–1.4
b11.9 b7.1% b13 b6.8
[8] [12] [23] [24]
29–102
b6.86 [26]
GC-PID HPLC-UV HPLC-UV HPLC-UV TD-GC–MS GC–MS
50–300 50–200 21–46 300–400 0.03–0.3 23.2–86.1
Solvent assisted DSPE Emulsification LPME DLLME Magnetic SPE SBSE Magnetically confined hydrophobic nanoparticles microextraction
Food and water River water Water Tap water Wastewater Wastewater Swimming pool and seawater samples
b8 b5.5 b9.3 b2.4 b13 b7.1
[27] [28] [29] [30] [31] This work
a Acronyms: SPME, solid phase microextraction; MEPS, microextraction by packed sorbent; SDME, single drop microextraction; LLE, liquid–liquid extraction; DLLME, dispersive liquid–liquid extraction; DSPE, dispersive solid phase extraction, LPME, liquid phase microextraction; SBSE, stir bar sorptive extraction. b Acronyms: GC, gas chromatography; MS, mass spectrometry; LVI, large volume injection; FID, flame ionization detector; PID, photo-ionization detector, HPLC, high performance liquid chromatography; UV, ultraviolet detection, TD, thermal desorption. c LOD, limit of detection. d RSD, relative standard deviation.
approach provides similar LODs than the obtained with dispersive liquid–liquid microextraction (DLLME) [26,29] and better results than those obtained with microextraction by packed sorbents (MEPS) [12], solvent assisted dispersive SPE [27], emulsification liquid phase microextraction [28] and magnetic SPE [30]. However, some of the latter approaches [12,30] have been successfully applied to the analysis of wastewater samples. Moreover, the rest of the methodologies [8,23,24,31] are more sensitive due to different reasons. The use of SPME [8] or SDME [23] involves a sensitivity enhancement since all the extracts (the fiber or the single drop) can be injected without a previous elution. On the other hand, the use of large volume injection (LVI) [24] or thermal desorption [31] allows the introduction of higher amounts of analytes in the instrument, improving also the sensitivity of the method. Concerning the precision, our method provides the fifth best value. Acknowledgments Financial support from the Spanish Ministry of Science and Innovation (grant CTQ2011-23790) and P09-FQM-4801 from the Junta de Andalucía are gratefully acknowledged. References [1] P.D. Darbre, A. Aljarrah, W.R. Miller, N.G. Coldham, M.J. Sauer, G.S. Pope, Concentrations of parabens in human breast tumours, J. Appl. Toxicol. 24 (2004) 5–13. [2] P.D. Darbre, Underarm cosmetics and breast cancer, J. Appl. Toxicol. 23 (2003) 89–95. [3] P.D. Darbre, P.W. Harvey, Paraben esters: review of recent studies of endocrine toxicity, absorption, esterase and human exposure, and discussion of potential human health risks, J. Appl. Toxicol. 28 (2008) 561–578. [4] E.J. Routledge, J. Parker, J. Odum, J. Ashby, J.P. Sumpter, Some alkyl hydroxyl benzoate preservatives (parabens) are estrogenic, Toxicol. Appl. Pharmacol. 153 (1998) 12–19. [5] R. Golden, J. Gandy, G. Vollmer, A review of the endocrine activity of parabens and implications for potential risks to human health, Crit. Rev. Toxicol. 35 (2005) 435–458. [6] J.M. Brausch, G.M. Rand, A review of personal care products in the aquatic environment: environmental concentrations and toxicity, Chemosphere 82 (2011) 1518–1532.
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