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Hollow fiber based liquid-phase microextraction for the determination of mercury traces in water samples by electrothermal atomic absorption spectrometry
Analytica Chimica Acta 743 (2012) 69–74
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Hollow fiber based liquid-phase microextraction for the determination of mercury traces in water samples by electrothermal atomic absorption spectrometry Ignacio López-García, Ricardo E. Rivas, Manuel Hernández-Córdoba ∗ Department of Analytical Chemistry, Faculty of Chemistry, Regional Campus of International Excellence “Campus Mare Nostrum”, University of Murcia, E-30071 Murcia, Spain
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
Hg (II) traces are preconcentrated by means of a three-phase liquid microextraction system. PAN and ammonium iodide are used in the donor and acceptor phase, respectively. Hollow-fiber pores are continuously fed with toluene placed in the lumen. Mercuric ions can be measured in waters below the g L−1 level.
a r t i c l e
i n f o
Article history: Received 18 May 2012 Received in revised form 10 July 2012 Accepted 11 July 2012 Available online 20 July 2012 Keywords: Liquid-phase microextraction Hollow fiber microextraction Mercury Electrothermal atomic absorption spectrometry Waters
a b s t r a c t A three-phase liquid microextraction procedure for the determination of mercury at low concentrations is discussed. To the aqueous sample placed at pH 7 by means of a phosphate buffer, 0.002% (m/v) 1-(2-pyridylazo)-2-naphthol (PAN) is incorporated, and the mixture submitted to microextraction with a hollow-fiber impregnated with toluene and whose lumen contains a 0.05 mol L−1 ammonium iodide solution. The final measurement of the extract is carried out by electrothermal atomic absorption spectrometry (300 ◦ C and 1100 ◦ C for the calcination and atomization temperatures, respectively). The pyrolytic graphite atomizer is coated electrolytically with palladium. An enrichment factor of 270, which results in a 0.06 g L−1 mercury for the detection limit is obtained. The relative standard deviation at the 1 g L−1 mercury level is 3.2% (n = 5). The reliability of the procedure is verified by analyzing waters as well as six certified reference materials. © 2012 Elsevier B.V. All rights reserved.
1. Introduction The success of solid-phase microextraction (SPME) has been the starting point for the development of new and ingenious ways of microextraction in the analytical laboratory. Following the pioneering works of Liu and Dasgupta [1] and Jeannot and Cantwell [2], who introduced liquid phase microextraction (LPME), a large number of approaches based on the distribution of the analytes between two immiscible liquids have been described [3–7]. One
∗ Corresponding author. Tel.: +34 868887406; fax: +34 868887682. E-mail address: [email protected] (M. Hernández-Córdoba). 0003-2670/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.aca.2012.07.015
of these microextraction techniques was proposed by PedersenBjergaard and Rasmussen and uses a hollow fiber (HF) to carry out the process (HF-LPME) [8]. In this approach, the HF acts as a support of an organic solvent which fills the lumen and is retained in the pores (two-phase operation). Alternatively, an aqueous solution can be placed inside the lumen of the HF, whose pores also contain an organic solvent. This results in a three-phase liquid microextraction, since the analytes are first transferred from the aqueous external solution (the sample or donor solution) toward the organic solvent microdrops inside the HF pores, and then into the aqueous inner solution (acceptor solution) in the lumen. In the beginning, HF-LPME was almost exclusively applied to the determination of organic compounds but, more recently, it
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Table 1 Instrumental parameters and furnace heating program. Parameter
Value
Lamp current (mA) Wavelength (nm) Spectral resolution (nm) Atomizer type Injected sample volume (L) Chemical modifier Background correction
6 253.7 0.5 Platform 10 Electrolytically reduced palladium coating Zeeman
Furnace program Stage
Temperature (◦ C)
Dry Ash Atomizationa
130 300 1100
a
Ramp (◦ C s−1 ) 10 10 0
Hold (s) 20 30 5
Reading stage.
has aroused increasing interest for the determination of very low concentrations of inorganic species due to the high enrichment factors that can be achieved [9–15]. Extensive work has been carried out by Hu et al. developing procedures for the speciation of selenium, vanadium and arsenic [16–18]. Using three-phase systems coupled to separation techniques, the speciation of organomercuric compounds has been also reported [19,20]. The same research group proposed the extraction of several diethyldithiocarbamatemetallic ion complexes [21]. The high enrichment factors even allow the use of classical molecular spectrophotometry in addition to atomic absorption spectrometry for the final measurement [22–25]. Another relevant manuscript deals with selenium determination [26] and, recently, a variant has been reported in which the organic solvent is replaced by an ion-pair [27]. In this work a procedure for the separation, preconcentration and determination of inorganic mercury traces with final measurement by electrothermal atomic absorption spectrometry (ETAAS) is optimized. The three-phase HF-LPME procedure achieves a high enrichment factor, which, together with the characteristics inherent in ETAAS measurements, results in a sensitive and selective procedure for the determination of very low concentrations of inorganic mercury. 2. Experimental 2.1. Instrumentation A model 939QZ atomic absorption spectrometer (ATI-Unicam, Cambridge, UK) equipped with both a deuterium and a Zeemanbased correction device, a longitudinally heated graphite furnace atomizer and pyrolytic graphite platforms inserted into pyrolytically coated graphite tubes were used. Argon was the inert gas, the flow rate being 300 mL min−1 during all the stages, except for atomization when the flow was stopped. Measurements were carried out in the peak-area mode using a mercury hollow cathode lamp operated at 6 mA and 253.7 nm with a 0.5 nm spectral bandwidth. The instrumental parameters used are summarized in Table 1. A microwave oven (Multiwave 3000 Anton-Paar, Perkin Elmer, Shelton, USA) provided with pressure and temperature control was used for the digestion of the reference materials. 2.2. Materials and reagents Ultrapure water, obtained using a Milli-Q system (Millipore, Bedford, MA, USA), was used exclusively. All glassware and plastic (polypropylene) vessels used for preparing and storing solutions were nitric acid-washed and rinsed with ultrapure water. Pipette tips were also of polypropylene. The Hg(II) stock standard
solution (1000 mg L−1 ) was obtained from Panreac (Barcelona, Spain). The rest of the chemicals used were obtained from Fluka (Buchs, Switzerland). Working standard Hg (II) solutions were prepared daily by appropriate dilution of the stock solution and incorporating 0.1% (v/v) concentrated nitric acid solution and 0.03 g L−1 potassium permanganate for stabilization purposes. A 10.0 g L−1 palladium solution (as nitrate) containing 15% (v/v) concentrated nitric acid solution was used for chemical modification as indicated below. The solutions of 1-(2-pyridylazo)-2-naphthol (PAN) were prepared in ethanol. Six certified reference materials were used to check the reliability of the procedure. Four of them, namely SRM 1571 (orchard leaves), SRM 1572 (citrus leaves), SRM 1515 (apple leaves) and SRM 1566a (oyster tissue), were obtained from the National Institute of Standards and Technology, while the others (dogfish muscle and dogfish liver, DORM-2 and DOLT-2, respectively) were provided by the National Research Council, Canada. The polypropylene hollow fibers (HF) were obtained from Membrana (Wuppertal, Germany). The specifications of the dealer indicate that these fibers have an internal diameter of 600 m and a 200 m wall thickness. The fibers, which are commercialized as 53 cm length strips, were cut into 16 cm segments, immersed in acetone and sonicated. After the evaporation of the organic liquid, the strips were ready to use as indicated below.
2.3. Treatment of the pyrolytic material Palladium was used as a permanent modifier. To this purpose, the pyrolytic material (both tube and platform) was first immersed overnight in a 10 g L−1 palladium solution. Next, the solution was electrolyzed for 3 min by applying 3 V between the graphite material, which acted as the cathode, and a small platinum wire as the anode. Finally, tube and platform were dried at 120 ◦ C for 15 min [28].
2.4. Procedure Two chromatographic syringes were used. The first (the main syringe) was used to handle the acceptor solution and the other, without the plunger, was used merely as a support to hold the polypropylene fiber. Firstly, a 30 mL sample aliquot was placed in a 50 mL vial, and 0.3 mL of 1 mol L−1 pH = 7 phosphate buffer solution and 0.3 mL of a 0.2% ethanolic PAN solution were incorporated. Then, 10 L of a 0.05 mol L−1 ammonium iodide solution and 10 L toluene were, in this order, drawn into the main syringe. A 16 cm HF strip was immersed for 10–15 s in toluene to facilitate the insertion of the 5 cm end sections into the tips of both syringes. The strip, in this position, was then again immersed in toluene for 25–30 s to impregnate the membrane pores before immersing in the vial containing the buffered sample solution. By pressing the plunger of the main syringe, the ammonium iodide acceptor solution was forced into the portion of the fiber (about 6 cm) which was in contact with the sample solution (donor solution). During the microextraction process (10 min), the solution was stirred magnetically. Once this time elapsed, the acceptor solution was drawn into the main syringe, the end of the strip connected to the auxiliary syringe was removed, the acceptor solution was transferred to the atomizer and the heating program described in Table 1 was run. Calibration was carried out against aqueous standard solutions submitted to the same procedure. The strips were discarded after each use. Graphical information as regards the arrangement of the syringes is provided as Fig. SI.1.
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Fig. 1. Optimization of the microextraction conditions. Toluene was used as the organic solvent. A 1.2 g L−1 Hg(II) solution was used. See the text for details.
2.5. Treatment of the standard reference materials The samples were digested by a microwave oven treatment. For this purpose, 3 mL concentrated nitric acid and 1 mL concentrated hydrogen peroxide were added to 0.1–0.3 g portions of the sample. The temperature-controlled digestion program consisted of a 10 min ramp from room temperature up to 180 ◦ C which was held for 10 min. After cooling, the acidity was neutralized with a 1 mol L−1 sodium hydroxide solution and the volume was finally made up to 25 mL. 3. Results and discussion 3.1. Optimization of the HF-LPME conditions For a metal to be transferred from an aqueous solution to an organic phase supported in a fiber, a complex must first be formed. Four complexing agents, namely ammonium pyrrolidine dithiocarbamate (APDC), 4-(2-pyridylazo)-resorcinol) (PAR), ammonium diethyl dithiophosphate (DDTP) and 1-(2-pyridylazo)-2-naphthol (PAN), were checked for the purpose, and PAN proved the most suitable. This classical reagent is a non-charged chemical in the 3–11 pH range, which forms complexes with a number of metallic ions. For the particular case of Hg (II), a 1:1 mercury:reagent complex was reported to be formed in aqueous media below pH 4, which changed to a 1:2 complex above pH 6 [29]. First at all, preliminary experiments were carried out, using toluene as the organic phase and a diluted iodide solution as the acceptor phase in the lumen of the HF. Next, the effect of the reagent concentration, acidity and pH on the extraction of the mercury–PAN complex into the acceptor solution were studied. The results, which are summarized in Fig. 1, allowed the optimal conditions to be selected. These were a PAN concentration of 0.002% (m/v) and pH 7, which was obtained by means of a phosphate buffer solution. The
final concentration of this buffer was raised to 0.01 mol L−1 to favor the salting-out effect without the need for additional salt [30]. An essential point in three-phase HF-LPME is the selection of the organic solvent to be retained in the pores of the HF. A suitable organic solvent for this purpose should have a low solubility in water and a high extraction coefficient for the complex originated in the aqueous sample. It is important to note that it is difficult to keep a few microliters of an organic solvent stable inside the HF pores. Since the extraction process lasts several minutes, and because the aqueous medium is being continuously stirred, the organic liquid acting as an intermediate is released from the HF pores to the aqueous donor solution, so that the process shows a poor reproducibility and can even stop. There are several ways to overcome this practical difficulty, the most simple probably involving a dispersion of the organic solvent in the aqueous phase [24,31]. However, poor reproducibility was found when this way was used in the case here studied. In contrast, reproducible results were obtained using a different approach based on a solvent reservoir. As described in Experimental, a microsyringe is first loaded with 10 L of acceptor aqueous liquid phase and next with 10 L of organic solvent. This low volume of organic liquid inside the lumen of the HF supplies fresh solvent to the pores, replacing the amount that is being released to the external aqueous donor solution. In this simple way, the HF pores are always fed with fresh solvent, and the microextraction process does not stop. Another relevant advantage is the decrease in the time required to carry out the microextraction. In practice, at the end of the process, the organic solvent inside the lumen is practically exhausted. Using this idea, several organic solvents in addition to toluene were assayed. Chloroform proved unsuitable due to its volatility, as occurred with cyclohexane and butyl acetate. On the contrary, for some solvents, such as iso-octane, the extraction was so efficient that it hindered the second extraction toward the aqueous solution placed in HF lumen. It was confirmed that toluene provided the best
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results. It is important to note that HF strips should not be reused to avoid cross-contamination. The time and reagents necessary for cleaning represent a cost that exceeds the low price of the HFs. In the liquid–liquid–liquid microextraction approach, the analyte is first transferred to the organic solvent in the HF pores, and then to the aqueous inner solution (acceptor solution) which is filling the HF lumen. For the case here discussed, the acceptor solution must contain a reagent forming water-soluble complexes with Hg(II), so that the Hg–PAN complex can be decomposed, the metallic ion passing to the internal solution. In this respect, and since the final measurement is carried out by ETAAS, the selection of this reagent is important to obtain a low background value during the atomization stage. A number of reagents, including cysteine, thiourea, ethylenediaminetetraacetate as well as chloride and iodide salts, were tried in an attempt to obtain an efficient extraction from toluene, while paying attention to the behavior of the reagent during the heating program of the atomizer. A 0.05 mol L−1 ammonium iodide solution (Fig. 1) was finally chosen. Sodium or potassium iodides proved inappropriate due to the high background values during electrothermal atomization, which rendered the final measurement difficult. As could be expected, both temperature and stirring of the sample solution affected the HF microextraction stage. An increase in the temperature would normally increase the rate of the process but the viscosity and water solubility of the organic liquid vary, reducing reproducibility. A study of the temperature was carried out in the 15–50 ◦ C range. No effect was noted below 30 ◦ C but above this value small bubbles were observed adhering to the external wall of the HF and both the analytical signal and the reproducibility decreased. It was concluded that no thermostatic control was required and all the experiments were carried out at room temperature. The stirring rate must be controlled because of the risk of forming a vortex in the vicinity of the HF immersed in the sample solution. For most of the experiments here reported, 29 mm external diameter vials containing 30 mL aqueous sample were used, in this way obtaining a solution column 50 mm high. With these dimensions, a small stirring bar (10 × 3 mm) rotating at 300 rpm provided efficient stirring without the formation of a vortex. Using the conditions thus optimized, the time required for the process was studied. Maximum and practically constant analytical signals were obtained when the microextraction lasted 10 min. Of note is the fact equilibrium was reached but mercury extraction to the HF lumen was incomplete, as confirmed by carrying out two consecutive extractions with the same aliquot of sample. Finally, in order to obtain the maximum analytical signal for mercury, a number of experiments were made varying the volume of the aqueous phase in the 10–50 mL range, while the volume of the acceptor solution filling the lumen was maintained at 10 L. A linear relationship between the analytical signal and the volume of the aqueous phase was found up to 35 mL, approximately. Consequently, 30 mL is recommended to obtain a high enrichment factor. On the other hand, since the literature reports [32] that PAN forms non-polar compounds with methylated species of mercury (methyl, ethyl and phenyl), additional experiments were devoted to checking whether these compounds passed into the acceptor iodide solution. No signal of mercury was obtained in the inner aqueous solutions even using 200 g L−1 of these compounds. The procedure here reported only allows inorganic Hg(II) to be measured. 3.2. Optimization of the ETAAS furnace program
Fig. 2. Ashing–atomizing graphs for mercury obtained in a 0.05 mol L−1 ammonium iodide solution and using the coating of reduced palladium for chemical modification.
of studies [28,33–42] have reported successful Hg-ETAAS measurements as long as a suitable chemical modifier is used in the procedure. From a practical point of view, it is advantageous to use permanent chemical modifiers rather than incorporating the modifier as an aqueous solution. With this in mind, a number of experiments were carried out in which the pyrolytic material was impregnated overnight by immersion in solutions containing gold, platinum, iridium or palladium salts. Additional experiments were carried out using electrolytic coating with palladium [28]. In all cases the performance of the treated atomizers for the ETAAS measurement of mercury was checked by atomizing aqueous solutions containing the metal both in the presence and absence of ammonium iodide buffered at pH 7, which is the composition of the acceptor solution inside the HF. As can be seen in Fig. 2, a calcination temperature of 300 ◦ C can be set when using electrolytic coating without a premature loss of analyte. Similar studies were carried out for the atomization temperature and 1100 ◦ C was found to be optimal. In these conditions (palladium electrolytic coating, 0.05 mol L−1 ammonium iodide at pH 7 and 300 ◦ C and 1100 ◦ C for calcination and atomization temperatures, respectively) welldefined atomization profiles for mercury were obtained (Fig. SI.2). The characteristic mass for mercury was found to be 63 pg, which is similar to that previously reported for this element in molten undecanoic acid [28]. Note that the chemical modification obtained with the electrolytic coating of palladium was effective for about 500 firings.
Table 2 Summary of the main figures of merit. Characteristic Linear range Hg(II) (g L−1 ) Detection limita Hg(II) (g L−1 ) Linear equationb Regression coefficient Relative standard deviationc (n = 5) (%) Sampling frequency (samples h−1 ) Sample volume (mL) Enrichment factord a
The determination of mercury by ETAAS is complicated by the high volatility of the element, which means that low temperatures should be used for the calcination stage. Despite this, a number
b c d
3sy/x criterium. Five standard solutions, n = 3. For a 1 g L−1 Hg(II) solution. Compared with the direct determination.
0.2–3 0.06 Aint = 0.008 + 0.1894CHg 0.9967 3.2 4 30 270
This work
[46]
[45]
[19] [44]
[21]
[43]
Ref.
I. López-García et al. / Analytica Chimica Acta 743 (2012) 69–74 Table 4 Results for the determination of mercury in different samples. Water samples
15
180
–
25 10
15
Bottled mineral water 2 –
Time (min)
Bottled mineral water 1
Bottled mineral water 3
0.06
0.8
Bottled mineral water 6 –
0.3–3.8 0.1
0.003
Bottled mineral water 5
–
LOD (g L−1 )
Bottled mineral water 4
Bottled mineral water 7
270
1000
Tap water 2
–
120, 215, 350 204
20
19
Tap water 1
EF
73
Tap water treated by reverse osmosis
Standard reference materials
Hg(II) (g L−1 ) Added
Founda
0 0.5 0 0.5 0 1.0 0 1.0 0 1.5 0 1.5 0 2.0 0 2.0 0 2.5 0 2.5
Hg content (g g−1 )
Visual test
0.01 M 2,6pyridinedicarboxylic acid 0.05 M NH4 I
pH = 2.5
2% NaCl, pH = 8
PAN, pH = 7
HgCl4 2−
Hg(II)
Hg(II)
Hg(II)
MeHg, EtHg, PhHg MeHg
2% (v/v) tri-n-octylamine in toluene 2% (m/v) trioctycphosphine oxide in undecane Toluene
ETAAS with Pd as chemical modifier
HPLC–UV ETAAS with Pd as chemical modifier Recovery and preconcentration 0.01 M Na2 S2 O3 4% thiourea, 1 M HCl Toluene Toluene
0.5 M NaOH
ICP-MS – CCl4
Recovery and preconcentration 0.3 M thiourea
0.1 M NaNO3 , pH = 2, N-benzoyl-N ,N diheptadecylthiourea 0.5% (m/v) diethyldithiocarbamate pH = 6 or 1 M HCl pH = 6 Hg(II)
80:20 decaline:cumene
Acceptor phase Extractant Donor phase Analyte
Table 3 Characteristics of same procedures reported for the HF-LPME preconcentration of mercury species.
Detection
Certified value SRM 1572 (citrus leaves) SRM 1515 (apple leaves) SRM 1566a (oyster tissue) NRC-CNRC DORM-2 (dogfish muscle) NRC-CNRC DOLT-2 (dogfish liver) a b
0.08 0.044 0.0642 4.64 2.14
± ± ± ± ±
0.02 0.004 0.0067 0.26 0.28
Foundb 0.07 0.04 0.06 4.56 2.18
± ± ± ± ±
0.02 0.01 0.02 0.12 0.10
Mean value (n = 5) ± standard deviation. Recovery percentage in brackets. Mean value (n = 5) ± standard deviation.
3.3. Analytical figures of merit: analysis of real samples The main figures of merit for the procedure optimized are shown in Table 2. The detection limit was calculated on the basis of three times the standard error from the calibrating regression (sy/x ). The enrichment factor was calculated as the ratio between the slope of a calibration graph obtained by submitting the samples to the microextraction process and the slope of another calibration graph obtained with samples that were not submitted to extraction but directly measured. The enrichment factor was found to be 270. Although this is a high value that allows a good sensitivity to be achieved, it confirms that mercury extraction is not total. The selectivity of the procedure is high due to the characteristics of the final ETAAS measurement. Other ions that react with PAN only interfere when present at relatively high concentrations due to the consumption of a significant fraction of PAN, and the subsequent decrease in the efficiency of the extraction. However, when present above 0.5 g L−1 chloride and bromide interfere since they hinder the formation of the Hg–PAN complex. Table 3 summarizes the procedures reported to date involving mercury and HF-LPME. Some of those mentioned were used exclusively for separation of the element in water for purification purposes. As can be seen, from several points of view, the comparison is favorable to the procedure here discussed. Finally, to check the reliability of the procedure, a number of real samples were analyzed, and the results are given in Table 4. As no mercury was found in any of the water samples analyzed, recovery studies were carried out. The practical usefulness of the procedure was confirmed by using six certified reference materials of different characteristics, and the results are also given in Table 4.
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4. Conclusion The procedure here presented shows good sensitivity, which is mainly a consequence of the high enrichment factor achieved in the microextraction process. The low detection limit (0.06 g L−1 ) allows the procedure to be used for the determination of mercury in potable waters since the limit established by the European Union is 1 g L−1 [47]. Taking into account that minimal amounts of commercial, relatively low cost reagents are involved, and the fact that ETAAS instruments are easily available in most laboratories, the procedure could be regarded as an alternative to those based on cold vapour generation. The short duration of the microextraction stage and the direct use of standard solutions for calibration without the need for standard additions are additional, positive features. In contrast, the discontinuous nature of both the microextraction stage and ETAAS measurement complicate automation. Acknowledgements The authors acknowledge to Comunidad Autónoma de la Región de Murcia (CARM, Fundación Séneca, Project 11796/PI/09) and to the Spanish MICINN (Project CTQ2009-08267/BQU) for financial support. R.E. Rivas acknowledges a fellowship of the Universidad CentroOccidental Lisandro Alvarado (UCLA), Venezuela. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.aca.2012.07.015. References [1] H. Liu, P.K. Dasgupta, Anal. Chem. 68 (1996) 1817–1821. [2] M.J. Jeannot, F.F. Cantwell, Anal. Chem. 68 (1996) 2236–2240. [3] A.L. Theis, A.J. Waldack, S.M. Hansen, M.A. Jeannot, Anal. Chem. 73 (2001) 5651–5654. [4] W. Liu, H.K. Lee, Anal. Chem. 72 (2000) 4462–4467. [5] l. Yangcheng, L. Quan, L. Guangsheng, D. Youyuan, Anal. Chim. Acta 566 (2006) 259–264. [6] M. Rezaee, Y. Assadi, M.R.M. Hosseini, E. Aghaee, F. Ahmadi, S. Berijani, J. Chromatogr. A 1116 (2006) 1–9. [7] H. Xu, Z. Ding, L. Lv, D. Song, Y.Q. Feng, Anal. Chim. Acta 636 (2009) 28–33. [8] S. Pedersen-Bjergaard, K.E. Rasmussen, Anal. Chem. 71 (1999) 2650–2656. [9] C. Nerín, J. Salafranca, M. Aznar, R. Batlle, Anal. Bioanal. Chem. 393 (2009) 809–833. [10] L. Xu, C. Basheer, H.K. Lee, J. Chromatogr. A 1216 (2009) 701–707.
[11] C.I.C. Silvestre, J.L.M. Santos, J.L.F.C. Lima, E.A.G. Zagatto, Anal. Chim. Acta 652 (2009) 54–65. [12] S. Dadfarnia, A.M.H. Shabani, Anal. Chim. Acta 658 (2010) 107–119. [13] F. Pena-Pereira, I. Lavilla, C. Bendicho, Anal. Chim. Acta 669 (2010) 1–16. [14] M. Cruz-Vera, R. Lucena, S. Cárdenas, M. Valcárcel, Anal. Methods 3 (2011) 1719–1728. [15] A. Jain, K.K. Verna, Anal. Chim. Acta 706 (2011) 37–65. [16] L. Xia, B. Hu, Z. Jiang, Y. Wu, R. Chen, L. Li, J. Anal. At. Spectrom. 21 (2006) 362–365. [17] L. Li, B. Hu, Talanta 72 (2007) 472–479. [18] H. Jiang, B. Hu, B. Chen, L. Xia, Anal. Chim. Acta 634 (2009) 15–21. [19] L. Xia, B. Hu, Y. Wu, J. Chromatogr. A 1173 (2007) 44–51. [20] P. Li, J. Duan, B. Hu, Electrophoresis 29 (2008) 3081–3089. [21] L. Xia, Y. Wu, B. Hu, J. Mass Spectrom. 42 (2007) 803–810. [22] R. Romero, J. Liu, P. Mayer, J.A. Jönsson, Anal. Chem. 77 (2005) 7605–7611. [23] J.S. Carletto, R.M. Luciano, G.C. Bedendo, E. Carasek, Anal. Chim. Acta 638 (2009) 45–50. [24] R.M. Luciano, G.C. Bedendo, J.S. Carletto, E. Carasek, J. Hazard. Mater. 177 (2010) 567–572. [25] J. Peng, R. Liu, J. Liu, B. He, X. Hu, G. Jiang, Spectrochim. Acta B 62 (2007) 499–503. [26] A. Saleh, Y. Yamini, M. Faraji, S. Shariati, M. Rezaee, J. Chromatogr. B 877 (2009) 1758–1764. [27] J. Abulhassani, J.L. Manzoori, M. Amjadi, J. Hazard. Mater. 176 (2010) 481–486. [28] I. López-García, R.E. Rivas, M. Hernández-Córdoba, Anal. Bioanal. Chem. 396 (2010) 3097–3102. [29] R.G. Anderson, G. Nickless, Analyst 92 (1967) 207–238. [30] E. Psillakis, N. Kalogerakis, Trac-Trends Anal. Chem. 22 (2003) 565–574. [31] D. Pezo, J. Salafranca, C. Nerín, J. Chromatogr. A 1174 (2007) 85–94. [32] X.B. Yin, J. Chromatogr. A 1154 (2007) 437–443. [33] A. Furtado da Silva, B. Welz, A.J. Curtius, Spectrochim. Acta B 57 (2002) 2031–2045. [34] P.R. Aranda, R.A. Gil, S. Moyano, I.E. De Vito, L.D. Martinez, Talanta 75 (2008) 307–311. [35] B. Welz, G. Schlemmer, J.R. Mudakavi, J. Anal. At. Spectrom. 7 (1992) 499–503. ˜ ˜ A. Moreda- Pineiro, A. Bermejo-Barrera, [36] P. Bermejo-Barrera, J. Moreda-Pineiro, Microchim. Acta 124 (1996) 111–122. [37] E. Bulska, W. Kandler, A. Hulanicki, Spectrochim. Acta B 51 (1996) 1263–1270. [38] I. López-García, M. Sánchez-Merlos, M. Hernández-Córdoba, Spectrochim. Acta B 52 (1997) 2085–2092. [39] W.N.L. Dos Santos, F.S. Dias, M.V. Reboucas, M.G. Pereira, V.A. Lemos, L.S.G. Teixeira, J. Anal. At. Spectrom. 21 (2006) 1327–1330. [40] P. Bermejo-Barrera, M. Aboal-Somoza, A. Bermejo-Barrera, M.L. Cervera, M. de la Guardia, J. Anal. At. Spectrom. 16 (2001) 382–389. [41] I. Karadjova, T. Venelinov, Food Add. Contam. 19 (2002) 948–953. [42] R.G.M. Moreno, E. de Oliveira, J.J. Pedrotti, P.V. Oliveira, Spectrochim. Acta B 67 (2002) 769–778. [43] C. Fontàs, M. Hidalgo, V. Salvadó, E. Anticó, Anal. Chim. Acta 547 (2005) 255–261. [44] H. Jiang, B. Hu, B. Chen, W. Zu, Spectrochim. Acta B 63 (2008) 770–776. [45] U. Pacharoen, S. Somboonpanya, S. Chatarabul, A.W. Lothongkum, J. Alloys Compd. 489 (2010) 72–79. [46] Z.Q. Tan, J.F. Liu, Anal. Chem. 82 (2010) 4222–4228. [47] Official Journal of the European Communities, COUNCIL DIRECTIVE 98/83/EC of 3 November 1998 on the quality of water intended for human consumption, L 330/32 (http://eur-lex.europa.eu/LexUriServ/ LexUriServ.do?uri=OJ:L:1998:330:0032:0054:EN:PDF).
Contents lists available at SciVerse ScienceDirect
Analytica Chimica Acta journal homepage: www.elsevier.com/locate/aca
Hollow fiber based liquid-phase microextraction for the determination of mercury traces in water samples by electrothermal atomic absorption spectrometry Ignacio López-García, Ricardo E. Rivas, Manuel Hernández-Córdoba ∗ Department of Analytical Chemistry, Faculty of Chemistry, Regional Campus of International Excellence “Campus Mare Nostrum”, University of Murcia, E-30071 Murcia, Spain
h i g h l i g h t s
g r a p h i c a l
a b s t r a c t
Hg (II) traces are preconcentrated by means of a three-phase liquid microextraction system. PAN and ammonium iodide are used in the donor and acceptor phase, respectively. Hollow-fiber pores are continuously fed with toluene placed in the lumen. Mercuric ions can be measured in waters below the g L−1 level.
a r t i c l e
i n f o
Article history: Received 18 May 2012 Received in revised form 10 July 2012 Accepted 11 July 2012 Available online 20 July 2012 Keywords: Liquid-phase microextraction Hollow fiber microextraction Mercury Electrothermal atomic absorption spectrometry Waters
a b s t r a c t A three-phase liquid microextraction procedure for the determination of mercury at low concentrations is discussed. To the aqueous sample placed at pH 7 by means of a phosphate buffer, 0.002% (m/v) 1-(2-pyridylazo)-2-naphthol (PAN) is incorporated, and the mixture submitted to microextraction with a hollow-fiber impregnated with toluene and whose lumen contains a 0.05 mol L−1 ammonium iodide solution. The final measurement of the extract is carried out by electrothermal atomic absorption spectrometry (300 ◦ C and 1100 ◦ C for the calcination and atomization temperatures, respectively). The pyrolytic graphite atomizer is coated electrolytically with palladium. An enrichment factor of 270, which results in a 0.06 g L−1 mercury for the detection limit is obtained. The relative standard deviation at the 1 g L−1 mercury level is 3.2% (n = 5). The reliability of the procedure is verified by analyzing waters as well as six certified reference materials. © 2012 Elsevier B.V. All rights reserved.
1. Introduction The success of solid-phase microextraction (SPME) has been the starting point for the development of new and ingenious ways of microextraction in the analytical laboratory. Following the pioneering works of Liu and Dasgupta [1] and Jeannot and Cantwell [2], who introduced liquid phase microextraction (LPME), a large number of approaches based on the distribution of the analytes between two immiscible liquids have been described [3–7]. One
∗ Corresponding author. Tel.: +34 868887406; fax: +34 868887682. E-mail address: [email protected] (M. Hernández-Córdoba). 0003-2670/$ – see front matter © 2012 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.aca.2012.07.015
of these microextraction techniques was proposed by PedersenBjergaard and Rasmussen and uses a hollow fiber (HF) to carry out the process (HF-LPME) [8]. In this approach, the HF acts as a support of an organic solvent which fills the lumen and is retained in the pores (two-phase operation). Alternatively, an aqueous solution can be placed inside the lumen of the HF, whose pores also contain an organic solvent. This results in a three-phase liquid microextraction, since the analytes are first transferred from the aqueous external solution (the sample or donor solution) toward the organic solvent microdrops inside the HF pores, and then into the aqueous inner solution (acceptor solution) in the lumen. In the beginning, HF-LPME was almost exclusively applied to the determination of organic compounds but, more recently, it
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Table 1 Instrumental parameters and furnace heating program. Parameter
Value
Lamp current (mA) Wavelength (nm) Spectral resolution (nm) Atomizer type Injected sample volume (L) Chemical modifier Background correction
6 253.7 0.5 Platform 10 Electrolytically reduced palladium coating Zeeman
Furnace program Stage
Temperature (◦ C)
Dry Ash Atomizationa
130 300 1100
a
Ramp (◦ C s−1 ) 10 10 0
Hold (s) 20 30 5
Reading stage.
has aroused increasing interest for the determination of very low concentrations of inorganic species due to the high enrichment factors that can be achieved [9–15]. Extensive work has been carried out by Hu et al. developing procedures for the speciation of selenium, vanadium and arsenic [16–18]. Using three-phase systems coupled to separation techniques, the speciation of organomercuric compounds has been also reported [19,20]. The same research group proposed the extraction of several diethyldithiocarbamatemetallic ion complexes [21]. The high enrichment factors even allow the use of classical molecular spectrophotometry in addition to atomic absorption spectrometry for the final measurement [22–25]. Another relevant manuscript deals with selenium determination [26] and, recently, a variant has been reported in which the organic solvent is replaced by an ion-pair [27]. In this work a procedure for the separation, preconcentration and determination of inorganic mercury traces with final measurement by electrothermal atomic absorption spectrometry (ETAAS) is optimized. The three-phase HF-LPME procedure achieves a high enrichment factor, which, together with the characteristics inherent in ETAAS measurements, results in a sensitive and selective procedure for the determination of very low concentrations of inorganic mercury. 2. Experimental 2.1. Instrumentation A model 939QZ atomic absorption spectrometer (ATI-Unicam, Cambridge, UK) equipped with both a deuterium and a Zeemanbased correction device, a longitudinally heated graphite furnace atomizer and pyrolytic graphite platforms inserted into pyrolytically coated graphite tubes were used. Argon was the inert gas, the flow rate being 300 mL min−1 during all the stages, except for atomization when the flow was stopped. Measurements were carried out in the peak-area mode using a mercury hollow cathode lamp operated at 6 mA and 253.7 nm with a 0.5 nm spectral bandwidth. The instrumental parameters used are summarized in Table 1. A microwave oven (Multiwave 3000 Anton-Paar, Perkin Elmer, Shelton, USA) provided with pressure and temperature control was used for the digestion of the reference materials. 2.2. Materials and reagents Ultrapure water, obtained using a Milli-Q system (Millipore, Bedford, MA, USA), was used exclusively. All glassware and plastic (polypropylene) vessels used for preparing and storing solutions were nitric acid-washed and rinsed with ultrapure water. Pipette tips were also of polypropylene. The Hg(II) stock standard
solution (1000 mg L−1 ) was obtained from Panreac (Barcelona, Spain). The rest of the chemicals used were obtained from Fluka (Buchs, Switzerland). Working standard Hg (II) solutions were prepared daily by appropriate dilution of the stock solution and incorporating 0.1% (v/v) concentrated nitric acid solution and 0.03 g L−1 potassium permanganate for stabilization purposes. A 10.0 g L−1 palladium solution (as nitrate) containing 15% (v/v) concentrated nitric acid solution was used for chemical modification as indicated below. The solutions of 1-(2-pyridylazo)-2-naphthol (PAN) were prepared in ethanol. Six certified reference materials were used to check the reliability of the procedure. Four of them, namely SRM 1571 (orchard leaves), SRM 1572 (citrus leaves), SRM 1515 (apple leaves) and SRM 1566a (oyster tissue), were obtained from the National Institute of Standards and Technology, while the others (dogfish muscle and dogfish liver, DORM-2 and DOLT-2, respectively) were provided by the National Research Council, Canada. The polypropylene hollow fibers (HF) were obtained from Membrana (Wuppertal, Germany). The specifications of the dealer indicate that these fibers have an internal diameter of 600 m and a 200 m wall thickness. The fibers, which are commercialized as 53 cm length strips, were cut into 16 cm segments, immersed in acetone and sonicated. After the evaporation of the organic liquid, the strips were ready to use as indicated below.
2.3. Treatment of the pyrolytic material Palladium was used as a permanent modifier. To this purpose, the pyrolytic material (both tube and platform) was first immersed overnight in a 10 g L−1 palladium solution. Next, the solution was electrolyzed for 3 min by applying 3 V between the graphite material, which acted as the cathode, and a small platinum wire as the anode. Finally, tube and platform were dried at 120 ◦ C for 15 min [28].
2.4. Procedure Two chromatographic syringes were used. The first (the main syringe) was used to handle the acceptor solution and the other, without the plunger, was used merely as a support to hold the polypropylene fiber. Firstly, a 30 mL sample aliquot was placed in a 50 mL vial, and 0.3 mL of 1 mol L−1 pH = 7 phosphate buffer solution and 0.3 mL of a 0.2% ethanolic PAN solution were incorporated. Then, 10 L of a 0.05 mol L−1 ammonium iodide solution and 10 L toluene were, in this order, drawn into the main syringe. A 16 cm HF strip was immersed for 10–15 s in toluene to facilitate the insertion of the 5 cm end sections into the tips of both syringes. The strip, in this position, was then again immersed in toluene for 25–30 s to impregnate the membrane pores before immersing in the vial containing the buffered sample solution. By pressing the plunger of the main syringe, the ammonium iodide acceptor solution was forced into the portion of the fiber (about 6 cm) which was in contact with the sample solution (donor solution). During the microextraction process (10 min), the solution was stirred magnetically. Once this time elapsed, the acceptor solution was drawn into the main syringe, the end of the strip connected to the auxiliary syringe was removed, the acceptor solution was transferred to the atomizer and the heating program described in Table 1 was run. Calibration was carried out against aqueous standard solutions submitted to the same procedure. The strips were discarded after each use. Graphical information as regards the arrangement of the syringes is provided as Fig. SI.1.
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Fig. 1. Optimization of the microextraction conditions. Toluene was used as the organic solvent. A 1.2 g L−1 Hg(II) solution was used. See the text for details.
2.5. Treatment of the standard reference materials The samples were digested by a microwave oven treatment. For this purpose, 3 mL concentrated nitric acid and 1 mL concentrated hydrogen peroxide were added to 0.1–0.3 g portions of the sample. The temperature-controlled digestion program consisted of a 10 min ramp from room temperature up to 180 ◦ C which was held for 10 min. After cooling, the acidity was neutralized with a 1 mol L−1 sodium hydroxide solution and the volume was finally made up to 25 mL. 3. Results and discussion 3.1. Optimization of the HF-LPME conditions For a metal to be transferred from an aqueous solution to an organic phase supported in a fiber, a complex must first be formed. Four complexing agents, namely ammonium pyrrolidine dithiocarbamate (APDC), 4-(2-pyridylazo)-resorcinol) (PAR), ammonium diethyl dithiophosphate (DDTP) and 1-(2-pyridylazo)-2-naphthol (PAN), were checked for the purpose, and PAN proved the most suitable. This classical reagent is a non-charged chemical in the 3–11 pH range, which forms complexes with a number of metallic ions. For the particular case of Hg (II), a 1:1 mercury:reagent complex was reported to be formed in aqueous media below pH 4, which changed to a 1:2 complex above pH 6 [29]. First at all, preliminary experiments were carried out, using toluene as the organic phase and a diluted iodide solution as the acceptor phase in the lumen of the HF. Next, the effect of the reagent concentration, acidity and pH on the extraction of the mercury–PAN complex into the acceptor solution were studied. The results, which are summarized in Fig. 1, allowed the optimal conditions to be selected. These were a PAN concentration of 0.002% (m/v) and pH 7, which was obtained by means of a phosphate buffer solution. The
final concentration of this buffer was raised to 0.01 mol L−1 to favor the salting-out effect without the need for additional salt [30]. An essential point in three-phase HF-LPME is the selection of the organic solvent to be retained in the pores of the HF. A suitable organic solvent for this purpose should have a low solubility in water and a high extraction coefficient for the complex originated in the aqueous sample. It is important to note that it is difficult to keep a few microliters of an organic solvent stable inside the HF pores. Since the extraction process lasts several minutes, and because the aqueous medium is being continuously stirred, the organic liquid acting as an intermediate is released from the HF pores to the aqueous donor solution, so that the process shows a poor reproducibility and can even stop. There are several ways to overcome this practical difficulty, the most simple probably involving a dispersion of the organic solvent in the aqueous phase [24,31]. However, poor reproducibility was found when this way was used in the case here studied. In contrast, reproducible results were obtained using a different approach based on a solvent reservoir. As described in Experimental, a microsyringe is first loaded with 10 L of acceptor aqueous liquid phase and next with 10 L of organic solvent. This low volume of organic liquid inside the lumen of the HF supplies fresh solvent to the pores, replacing the amount that is being released to the external aqueous donor solution. In this simple way, the HF pores are always fed with fresh solvent, and the microextraction process does not stop. Another relevant advantage is the decrease in the time required to carry out the microextraction. In practice, at the end of the process, the organic solvent inside the lumen is practically exhausted. Using this idea, several organic solvents in addition to toluene were assayed. Chloroform proved unsuitable due to its volatility, as occurred with cyclohexane and butyl acetate. On the contrary, for some solvents, such as iso-octane, the extraction was so efficient that it hindered the second extraction toward the aqueous solution placed in HF lumen. It was confirmed that toluene provided the best
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results. It is important to note that HF strips should not be reused to avoid cross-contamination. The time and reagents necessary for cleaning represent a cost that exceeds the low price of the HFs. In the liquid–liquid–liquid microextraction approach, the analyte is first transferred to the organic solvent in the HF pores, and then to the aqueous inner solution (acceptor solution) which is filling the HF lumen. For the case here discussed, the acceptor solution must contain a reagent forming water-soluble complexes with Hg(II), so that the Hg–PAN complex can be decomposed, the metallic ion passing to the internal solution. In this respect, and since the final measurement is carried out by ETAAS, the selection of this reagent is important to obtain a low background value during the atomization stage. A number of reagents, including cysteine, thiourea, ethylenediaminetetraacetate as well as chloride and iodide salts, were tried in an attempt to obtain an efficient extraction from toluene, while paying attention to the behavior of the reagent during the heating program of the atomizer. A 0.05 mol L−1 ammonium iodide solution (Fig. 1) was finally chosen. Sodium or potassium iodides proved inappropriate due to the high background values during electrothermal atomization, which rendered the final measurement difficult. As could be expected, both temperature and stirring of the sample solution affected the HF microextraction stage. An increase in the temperature would normally increase the rate of the process but the viscosity and water solubility of the organic liquid vary, reducing reproducibility. A study of the temperature was carried out in the 15–50 ◦ C range. No effect was noted below 30 ◦ C but above this value small bubbles were observed adhering to the external wall of the HF and both the analytical signal and the reproducibility decreased. It was concluded that no thermostatic control was required and all the experiments were carried out at room temperature. The stirring rate must be controlled because of the risk of forming a vortex in the vicinity of the HF immersed in the sample solution. For most of the experiments here reported, 29 mm external diameter vials containing 30 mL aqueous sample were used, in this way obtaining a solution column 50 mm high. With these dimensions, a small stirring bar (10 × 3 mm) rotating at 300 rpm provided efficient stirring without the formation of a vortex. Using the conditions thus optimized, the time required for the process was studied. Maximum and practically constant analytical signals were obtained when the microextraction lasted 10 min. Of note is the fact equilibrium was reached but mercury extraction to the HF lumen was incomplete, as confirmed by carrying out two consecutive extractions with the same aliquot of sample. Finally, in order to obtain the maximum analytical signal for mercury, a number of experiments were made varying the volume of the aqueous phase in the 10–50 mL range, while the volume of the acceptor solution filling the lumen was maintained at 10 L. A linear relationship between the analytical signal and the volume of the aqueous phase was found up to 35 mL, approximately. Consequently, 30 mL is recommended to obtain a high enrichment factor. On the other hand, since the literature reports [32] that PAN forms non-polar compounds with methylated species of mercury (methyl, ethyl and phenyl), additional experiments were devoted to checking whether these compounds passed into the acceptor iodide solution. No signal of mercury was obtained in the inner aqueous solutions even using 200 g L−1 of these compounds. The procedure here reported only allows inorganic Hg(II) to be measured. 3.2. Optimization of the ETAAS furnace program
Fig. 2. Ashing–atomizing graphs for mercury obtained in a 0.05 mol L−1 ammonium iodide solution and using the coating of reduced palladium for chemical modification.
of studies [28,33–42] have reported successful Hg-ETAAS measurements as long as a suitable chemical modifier is used in the procedure. From a practical point of view, it is advantageous to use permanent chemical modifiers rather than incorporating the modifier as an aqueous solution. With this in mind, a number of experiments were carried out in which the pyrolytic material was impregnated overnight by immersion in solutions containing gold, platinum, iridium or palladium salts. Additional experiments were carried out using electrolytic coating with palladium [28]. In all cases the performance of the treated atomizers for the ETAAS measurement of mercury was checked by atomizing aqueous solutions containing the metal both in the presence and absence of ammonium iodide buffered at pH 7, which is the composition of the acceptor solution inside the HF. As can be seen in Fig. 2, a calcination temperature of 300 ◦ C can be set when using electrolytic coating without a premature loss of analyte. Similar studies were carried out for the atomization temperature and 1100 ◦ C was found to be optimal. In these conditions (palladium electrolytic coating, 0.05 mol L−1 ammonium iodide at pH 7 and 300 ◦ C and 1100 ◦ C for calcination and atomization temperatures, respectively) welldefined atomization profiles for mercury were obtained (Fig. SI.2). The characteristic mass for mercury was found to be 63 pg, which is similar to that previously reported for this element in molten undecanoic acid [28]. Note that the chemical modification obtained with the electrolytic coating of palladium was effective for about 500 firings.
Table 2 Summary of the main figures of merit. Characteristic Linear range Hg(II) (g L−1 ) Detection limita Hg(II) (g L−1 ) Linear equationb Regression coefficient Relative standard deviationc (n = 5) (%) Sampling frequency (samples h−1 ) Sample volume (mL) Enrichment factord a
The determination of mercury by ETAAS is complicated by the high volatility of the element, which means that low temperatures should be used for the calcination stage. Despite this, a number
b c d
3sy/x criterium. Five standard solutions, n = 3. For a 1 g L−1 Hg(II) solution. Compared with the direct determination.
0.2–3 0.06 Aint = 0.008 + 0.1894CHg 0.9967 3.2 4 30 270
This work
[46]
[45]
[19] [44]
[21]
[43]
Ref.
I. López-García et al. / Analytica Chimica Acta 743 (2012) 69–74 Table 4 Results for the determination of mercury in different samples. Water samples
15
180
–
25 10
15
Bottled mineral water 2 –
Time (min)
Bottled mineral water 1
Bottled mineral water 3
0.06
0.8
Bottled mineral water 6 –
0.3–3.8 0.1
0.003
Bottled mineral water 5
–
LOD (g L−1 )
Bottled mineral water 4
Bottled mineral water 7
270
1000
Tap water 2
–
120, 215, 350 204
20
19
Tap water 1
EF
73
Tap water treated by reverse osmosis
Standard reference materials
Hg(II) (g L−1 ) Added
Founda
0 0.5 0 0.5 0 1.0 0 1.0 0 1.5 0 1.5 0 2.0 0 2.0 0 2.5 0 2.5
Hg content (g g−1 )
Visual test
0.01 M 2,6pyridinedicarboxylic acid 0.05 M NH4 I
pH = 2.5
2% NaCl, pH = 8
PAN, pH = 7
HgCl4 2−
Hg(II)
Hg(II)
Hg(II)
MeHg, EtHg, PhHg MeHg
2% (v/v) tri-n-octylamine in toluene 2% (m/v) trioctycphosphine oxide in undecane Toluene
ETAAS with Pd as chemical modifier
HPLC–UV ETAAS with Pd as chemical modifier Recovery and preconcentration 0.01 M Na2 S2 O3 4% thiourea, 1 M HCl Toluene Toluene
0.5 M NaOH
ICP-MS – CCl4
Recovery and preconcentration 0.3 M thiourea
0.1 M NaNO3 , pH = 2, N-benzoyl-N ,N diheptadecylthiourea 0.5% (m/v) diethyldithiocarbamate pH = 6 or 1 M HCl pH = 6 Hg(II)
80:20 decaline:cumene
Acceptor phase Extractant Donor phase Analyte
Table 3 Characteristics of same procedures reported for the HF-LPME preconcentration of mercury species.
Detection
Certified value SRM 1572 (citrus leaves) SRM 1515 (apple leaves) SRM 1566a (oyster tissue) NRC-CNRC DORM-2 (dogfish muscle) NRC-CNRC DOLT-2 (dogfish liver) a b
0.08 0.044 0.0642 4.64 2.14
± ± ± ± ±
0.02 0.004 0.0067 0.26 0.28
Foundb 0.07 0.04 0.06 4.56 2.18
± ± ± ± ±
0.02 0.01 0.02 0.12 0.10
Mean value (n = 5) ± standard deviation. Recovery percentage in brackets. Mean value (n = 5) ± standard deviation.
3.3. Analytical figures of merit: analysis of real samples The main figures of merit for the procedure optimized are shown in Table 2. The detection limit was calculated on the basis of three times the standard error from the calibrating regression (sy/x ). The enrichment factor was calculated as the ratio between the slope of a calibration graph obtained by submitting the samples to the microextraction process and the slope of another calibration graph obtained with samples that were not submitted to extraction but directly measured. The enrichment factor was found to be 270. Although this is a high value that allows a good sensitivity to be achieved, it confirms that mercury extraction is not total. The selectivity of the procedure is high due to the characteristics of the final ETAAS measurement. Other ions that react with PAN only interfere when present at relatively high concentrations due to the consumption of a significant fraction of PAN, and the subsequent decrease in the efficiency of the extraction. However, when present above 0.5 g L−1 chloride and bromide interfere since they hinder the formation of the Hg–PAN complex. Table 3 summarizes the procedures reported to date involving mercury and HF-LPME. Some of those mentioned were used exclusively for separation of the element in water for purification purposes. As can be seen, from several points of view, the comparison is favorable to the procedure here discussed. Finally, to check the reliability of the procedure, a number of real samples were analyzed, and the results are given in Table 4. As no mercury was found in any of the water samples analyzed, recovery studies were carried out. The practical usefulness of the procedure was confirmed by using six certified reference materials of different characteristics, and the results are also given in Table 4.
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4. Conclusion The procedure here presented shows good sensitivity, which is mainly a consequence of the high enrichment factor achieved in the microextraction process. The low detection limit (0.06 g L−1 ) allows the procedure to be used for the determination of mercury in potable waters since the limit established by the European Union is 1 g L−1 [47]. Taking into account that minimal amounts of commercial, relatively low cost reagents are involved, and the fact that ETAAS instruments are easily available in most laboratories, the procedure could be regarded as an alternative to those based on cold vapour generation. The short duration of the microextraction stage and the direct use of standard solutions for calibration without the need for standard additions are additional, positive features. In contrast, the discontinuous nature of both the microextraction stage and ETAAS measurement complicate automation. Acknowledgements The authors acknowledge to Comunidad Autónoma de la Región de Murcia (CARM, Fundación Séneca, Project 11796/PI/09) and to the Spanish MICINN (Project CTQ2009-08267/BQU) for financial support. R.E. Rivas acknowledges a fellowship of the Universidad CentroOccidental Lisandro Alvarado (UCLA), Venezuela. Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.aca.2012.07.015. References [1] H. Liu, P.K. Dasgupta, Anal. Chem. 68 (1996) 1817–1821. [2] M.J. Jeannot, F.F. Cantwell, Anal. Chem. 68 (1996) 2236–2240. [3] A.L. Theis, A.J. Waldack, S.M. Hansen, M.A. Jeannot, Anal. Chem. 73 (2001) 5651–5654. [4] W. Liu, H.K. Lee, Anal. Chem. 72 (2000) 4462–4467. [5] l. Yangcheng, L. Quan, L. Guangsheng, D. Youyuan, Anal. Chim. Acta 566 (2006) 259–264. [6] M. Rezaee, Y. Assadi, M.R.M. Hosseini, E. Aghaee, F. Ahmadi, S. Berijani, J. Chromatogr. A 1116 (2006) 1–9. [7] H. Xu, Z. Ding, L. Lv, D. Song, Y.Q. Feng, Anal. Chim. Acta 636 (2009) 28–33. [8] S. Pedersen-Bjergaard, K.E. Rasmussen, Anal. Chem. 71 (1999) 2650–2656. [9] C. Nerín, J. Salafranca, M. Aznar, R. Batlle, Anal. Bioanal. Chem. 393 (2009) 809–833. [10] L. Xu, C. Basheer, H.K. Lee, J. Chromatogr. A 1216 (2009) 701–707.
[11] C.I.C. Silvestre, J.L.M. Santos, J.L.F.C. Lima, E.A.G. Zagatto, Anal. Chim. Acta 652 (2009) 54–65. [12] S. Dadfarnia, A.M.H. Shabani, Anal. Chim. Acta 658 (2010) 107–119. [13] F. Pena-Pereira, I. Lavilla, C. Bendicho, Anal. Chim. Acta 669 (2010) 1–16. [14] M. Cruz-Vera, R. Lucena, S. Cárdenas, M. Valcárcel, Anal. Methods 3 (2011) 1719–1728. [15] A. Jain, K.K. Verna, Anal. Chim. Acta 706 (2011) 37–65. [16] L. Xia, B. Hu, Z. Jiang, Y. Wu, R. Chen, L. Li, J. Anal. At. Spectrom. 21 (2006) 362–365. [17] L. Li, B. Hu, Talanta 72 (2007) 472–479. [18] H. Jiang, B. Hu, B. Chen, L. Xia, Anal. Chim. Acta 634 (2009) 15–21. [19] L. Xia, B. Hu, Y. Wu, J. Chromatogr. A 1173 (2007) 44–51. [20] P. Li, J. Duan, B. Hu, Electrophoresis 29 (2008) 3081–3089. [21] L. Xia, Y. Wu, B. Hu, J. Mass Spectrom. 42 (2007) 803–810. [22] R. Romero, J. Liu, P. Mayer, J.A. Jönsson, Anal. Chem. 77 (2005) 7605–7611. [23] J.S. Carletto, R.M. Luciano, G.C. Bedendo, E. Carasek, Anal. Chim. Acta 638 (2009) 45–50. [24] R.M. Luciano, G.C. Bedendo, J.S. Carletto, E. Carasek, J. Hazard. Mater. 177 (2010) 567–572. [25] J. Peng, R. Liu, J. Liu, B. He, X. Hu, G. Jiang, Spectrochim. Acta B 62 (2007) 499–503. [26] A. Saleh, Y. Yamini, M. Faraji, S. Shariati, M. Rezaee, J. Chromatogr. B 877 (2009) 1758–1764. [27] J. Abulhassani, J.L. Manzoori, M. Amjadi, J. Hazard. Mater. 176 (2010) 481–486. [28] I. López-García, R.E. Rivas, M. Hernández-Córdoba, Anal. Bioanal. Chem. 396 (2010) 3097–3102. [29] R.G. Anderson, G. Nickless, Analyst 92 (1967) 207–238. [30] E. Psillakis, N. Kalogerakis, Trac-Trends Anal. Chem. 22 (2003) 565–574. [31] D. Pezo, J. Salafranca, C. Nerín, J. Chromatogr. A 1174 (2007) 85–94. [32] X.B. Yin, J. Chromatogr. A 1154 (2007) 437–443. [33] A. Furtado da Silva, B. Welz, A.J. Curtius, Spectrochim. Acta B 57 (2002) 2031–2045. [34] P.R. Aranda, R.A. Gil, S. Moyano, I.E. De Vito, L.D. Martinez, Talanta 75 (2008) 307–311. [35] B. Welz, G. Schlemmer, J.R. Mudakavi, J. Anal. At. Spectrom. 7 (1992) 499–503. ˜ ˜ A. Moreda- Pineiro, A. Bermejo-Barrera, [36] P. Bermejo-Barrera, J. Moreda-Pineiro, Microchim. Acta 124 (1996) 111–122. [37] E. Bulska, W. Kandler, A. Hulanicki, Spectrochim. Acta B 51 (1996) 1263–1270. [38] I. López-García, M. Sánchez-Merlos, M. Hernández-Córdoba, Spectrochim. Acta B 52 (1997) 2085–2092. [39] W.N.L. Dos Santos, F.S. Dias, M.V. Reboucas, M.G. Pereira, V.A. Lemos, L.S.G. Teixeira, J. Anal. At. Spectrom. 21 (2006) 1327–1330. [40] P. Bermejo-Barrera, M. Aboal-Somoza, A. Bermejo-Barrera, M.L. Cervera, M. de la Guardia, J. Anal. At. Spectrom. 16 (2001) 382–389. [41] I. Karadjova, T. Venelinov, Food Add. Contam. 19 (2002) 948–953. [42] R.G.M. Moreno, E. de Oliveira, J.J. Pedrotti, P.V. Oliveira, Spectrochim. Acta B 67 (2002) 769–778. [43] C. Fontàs, M. Hidalgo, V. Salvadó, E. Anticó, Anal. Chim. Acta 547 (2005) 255–261. [44] H. Jiang, B. Hu, B. Chen, W. Zu, Spectrochim. Acta B 63 (2008) 770–776. [45] U. Pacharoen, S. Somboonpanya, S. Chatarabul, A.W. Lothongkum, J. Alloys Compd. 489 (2010) 72–79. [46] Z.Q. Tan, J.F. Liu, Anal. Chem. 82 (2010) 4222–4228. [47] Official Journal of the European Communities, COUNCIL DIRECTIVE 98/83/EC of 3 November 1998 on the quality of water intended for human consumption, L 330/32 (http://eur-lex.europa.eu/LexUriServ/ LexUriServ.do?uri=OJ:L:1998:330:0032:0054:EN:PDF).