Simultaneous Determination of Estrogenic Short Ethoxy Chain

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Anal. Chem. 2002, 74, 3869-3876

Simultaneous Determination of Estrogenic Short Ethoxy Chain Nonylphenols and Their Acidic Metabolites in Water by an In-Sample Derivatization/Solid-Phase Microextraction Method Alfredo Dı´az and Francesc Ventura*

AGBAR, Aigu¨es de Barcelona, P. Sant Joan, 39, 08009-Barcelona, Spain Maria Teresa Galceran

Department of Analytical Chemistry, University of Barcelona, Martı´ i Franque` s, 1-11, 08028-Barcelona, Spain

An in-sample derivatization headspace solid-phase microextraction method has been developed for the simultaneous determination of nonylphenol, nonylphenol monoand diethoxylates (NP, NP1EO, NP2EO), and their acidic metabolites (NP1EC, NP2EC) in water. The analytical procedure involves derivatization of NPEOs and NPECs to their methyl ethers-esters with dimethyl sulfate/NaOH and further headspace (HS) solid-phase microextraction (SPME) and gas chromatography/mass spectrometry (GC/MS) determination. Parameters affecting both derivatization efficiency and headspace SPME procedure, such as the selection of the SPME coating, derivatizationextraction time, temperature and ionic strength were optimized. The commercially available Carbowax-divinylbenzene (CW-DVB) fiber appears to be the most suitable for the simultaneous determination of both NPEOs-NPECs. Run-to-run precision of the in-sample derivatization/HS-SPME-GC/MS method gave relative standard deviations between 8 and 18%. The method was linear for NP over 2 orders of magnitude, and detection limits were compound dependent but ranged from 20 to 1500 ng/L. The SPME procedure was compared with a solid-phase extraction SPE-GC/MS method for the analysis of NPEOs-NPECs in water samples and good agreement was obtained. Therefore, in-sample derivatization HS-SPME-GC/MS can be used as a method for the simultaneous determination of short ethoxy chain nonylphenols and their acidic metabolites in water. Nonionic surfactants account for roughly 40% of worldwide surfactant market. Among them, alkylphenol polyethoxylates (APnEOs, n ) number of ethoxy units), with a world annual production rate of 500 000 tons in 1996, are widely used in industrial formulations (textile, tannery, paper industries, metal working fluids), pesticide adjuvants, paint ingredients, and personal care products.1 Approximately 80% of the APnEOs are * Corresponding author: (e-mail) [email protected]; (fax) +34 93 342 3666. (1) Renner, R. Environ. Sci. Technol. 1997, 31, 316A-320A. 10.1021/ac020124p CCC: $22.00 Published on Web 07/04/2002

© 2002 American Chemical Society

Figure 1. Acronyms and structures of the studied compounds.

nonylphenol ethoxylates (NPEOs) while the remaining 20% are mainly octylphenol ethoxylates (OPnEOs). The biodegradation of the parent compound (i.e., NPnEO, n ≈ 20-30), by progressive shortening of the ethoxylate chain under aerobic conditions, leads mainly to the formation of nonylphenol mono- and diethoxylates,2,3 whereas under anaerobic conditions, fully deethoxylated nonylphenol is also produced.4 Further transformation proceeds via oxidation of the ethoxylate chain producing nonylphenoxyacetic acid (NP1EC) and nonylphenoxyethoxyacetic acid (NP2EC). Figure 1 displays the chemical structures and acronyms used in this work. Recently, the presence of dicarboxylated metabolites of NPEOs in treated sewage sludge,5 and river water,6 has also been reported. In this case,6 the authors proposed an ω-carboxylation of the ethoxylate chain as initiating step of the biodegradation and a further transformation into short-chain ethoxy carboxylates (NPECs), with the nonylphenoxyethoxyacetic acid being the most abundant species. (2) Ahel, M.; Giger, W.; Schaffner, C. Water Res. 1994, 28, 1143-1152. (3) Ahel, M.; Giger, W. Water Res. 1994, 28, 1131-1142. (4) Giger, W.; Brunner, P. H.; Schaffner, C. Science 1984, 225, 623-625. (5) Di Corcia, A.; Cavallo, R.; Crescenzi, C.; Nazzari, M. Environ. Sci. Technol. 2000, 34, 3914-3919. (6) Jonkers, N.; Knepper, T. P.; De Voogt, P. Environ. Sci. Technol. 2001, 35, 335-340.

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Concern about NP and short ethoxy chain metabolites that exhibit toxicity to several aquatic organisms,7 and can bioaccumulate in aquatic food chains,8-10 has been raised due to their lipophilic properties (log Kow > 4). Moreover, several reports have demonstrated that these compounds are endocrine disruptors and can cause estrogenic effects in fish and other organisms.11-13 Different analytical methods have been proposed to determine these contaminants in different matrixes.14,15 Some of them used gas chromatography coupled with mass spectrometry (GC/MS), although this technique is restricted to the analysis of short ethoxy chain metabolites because of the limited volatility of the high molecular weight homologues.16-19 Liquid chromatography (LC) with UV and fluorescence detection (FD) has been extensively employed to determine both, parent (NPEO) and their metabolites,20-26 but nowadays liquid chromatography coupled to mass spectrometry (LC/MS) with electrospray ionization is increasingly used.27-30 The extraction and preconcentration methods of the analytes most frequently employed are solvent sublation,22 steam distillation,20,31 liquid-liquid extraction,32 and solid-phase extraction.24,25 Solid-phase microextraction (SPME) pionereed by Pawliszyn and co-workers,33-35 has been successfully applied for a wide range of organic compounds in water samples.36-38 However, there are very few papers dealing with the use of this technique for the (7) Servos, M. R. Water Qual. Res. J. Can. 1999, 34, 123-177. (8) Ahel, M.; Giger, W. Chemosphere 1993, 26, 1470-1478. (9) Ekelund, R.; Hergman, A.; Granmo, A.; Berggren, M. Environ. Pollut. 1990, 64, 107-120. (10) Ahel, M.; Giger, W.; McEvoy, J. Environ. Pollut. 1993, 79, 243-248. (11) Soto, A. M., Justicia, H.; Wary, W. J.; Sonnenschein, C. Environ. Health Perspect. 1991, 92,167-173. (12) Jobling, S.; Sumpter, J. P. Aquat. Toxicol. 1993, 27, 361-372. (13) Jobling, S.; Sheahan, D.; Osborne, J. A.; Matthiessen, P.; Sumpter, J. P. Environ. Toxicol. Chem. 1996, 15, 194-202. (14) Thiele, B.; Gu ¨ nther, K.; Schwuger, M. J. Chem. Rev. 1997, 97, 3247-3272. (15) Lee, H. B. Water Qual. Res. J. Can. 1999, 34, 3-35. (16) Stephanou, E.; Giger, W. Environ. Sci. Technol. 1982, 16, 800-805. (17) Reinhard, M.; Goodman, N.; Mortelmans, K. E. Environ. Sci. Technol. 1982, 16, 351-362. (18) Ventura, F.; Figueras, A.; Caixach, J.; Espadaler, I.; Romero, J.; Guardiola, J.; Rivera, J. Water Res. 1988, 22, 1211-1217. (19) Field, J. A.; Reed, R. L. Environ. Sci. Technol. 1996, 30, 3544-3550. (20) Ahel, M.; Giger, W. Anal. Chem. 1985, 57, 1577-1583. (21) Marcomini, A.; Giger, W. Anal. Chem. 1987, 59, 1709-1715. (22) Ahel, M.; Giger, W. Anal. Chem. 1985, 57, 2584-2590. (23) Marcomini, A.; Capri, S.; Giger, W. J. Chromatogr., A 1987, 403, 243-252. (24) Di Corcia, A.; Samperi, R.; Marcomini, A. Environ. Sci. Technol. 1991, 28, 850-858. (25) Crescenzi, C.; Di Corcia, A.; Samperi, R.; Marcomini, A. Anal. Chem. 1995, 67, 1797-1804. (26) Ahel, M.; Giger, W.; Molnar, E.; Ibric, S. Croatia Chem. Acta 2000, 73, 209-227. (27) Crescenzi, C.; Di Corcia, A.; Samperi, R.; Marcomini, A. Anal. Chem. 1995, 67, 1797-1804. (28) Di Corcia. J. Chromatogr., A 1998, 794, 165-185. (29) Cathum, S.; Sabik, H. Chromatographia 2001, 53, S-400-404. (30) Petrovic, M.; Diaz, A.; Ventura, F.; Barcelo´, D. Anal. Chem. 2001, 73, 58865895. (31) Giger, W.; Stephanou, E.; Schaffner, C. Chemosphere 1981, 10, 1253-1263. (32) Ahel, M.; Conrad, T.; Giger, W. Environ. Sci. Technol. 1987, 21, 697-703. (33) Arthur, C. L.; Killiam, L. M.; Buchholz, K. D.; Pawliszyn, J. Anal. Chem. 1992, 64, 1960-1966. (34) Zhang, Z.; Pawliszyn, J. Anal. Chem. 1993, 65, 1843-1852. (35) Pawliszyn, J. Solid-Phase Microextraction: Theory and Practice; Wiley-VCH: New York, 1997. (36) Pawliszyn, J., Ed. Applications of Solid-Phase Microextraction; RSC Chromatography Monographs; Royal Society of Chemistry: Cambridge, U.K., 1999. (37) Nilsson, T.; Pelusio, F.; Montanarella, L.; Larsen, B.; Facchetti, S.; Madsen, J. O. J. High Resolut. Chromatogr. 1995, 18, 617-624. (38) Eisert, R.; Levsen, K. J. Chromatogr., A 1996, 733, 143-157.

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determination of surfactants in water. The use of SPME-LC-FD for the analysis of anionic surfactants, such as linear alkylbenzenesulfonates (LAS) in influent and effluent wastewater samples of a sewage wastewater plant (STP), has been studied but the method was unsuitable to quantify LAS because of its limited extraction efficiency.39 Oxidation products of anionic surfactants such as lauryl sulfate and fatty alcohol polyethoxy sulfates have been identified by SPME-GC/MS and also by SPME-LC/MS.40,41 The first paper dealing with the determination of 4-nonylphenol in water by SPME used a poly(dimethylsiloxane) (PDMS) fiber and analysis by gas chromatography (GC-FID). With this method, an estimated detection limit of 0.1 mg/L42 was obtained. For the analysis of nonionic surfactants (APnEOs), Boyd-Boland and Pawliszyn,43 used SPME-LC-UV with normal-phase gradient elution. Different fibers were tested, and the best results were obtained with the Carbowax-template resin (CW-TR) fiber, which provided the best agreement between the distribution of ethoxamers before and after extraction and a limit of detection for individual alkylphenol ethoxamers at the low-microgram per liter level. The recent development of new coatings has expanded the range of compounds that can be analyzed by SPME. Thus, the determination of short ethoxy chain metabolites of nonylphenols and their halogenated derivatives formed by chlorination practices in water treatment plants using SPME has been proposed.44 Other nonionic surfactants such as alcohol polyethoxylates have been determined by SPME-LC-FD using a divinylbenzene (DVB)-Carboxen-PDMS fiber and precolumn derivatization with 1-naphthoyl chloride45 and a detection limit of 0.1 mg/L in water was obtained. The aim of this work is to develop a method for the simultaneous analysis of the estrogenic short ethoxy chain nonylphenol ethoxylates and their acidic metabolites produced in the biodegradation of NPEOs. Two different approaches involving on-fiber derivatization aqueous-SPME and in-sample derivatization headspace-SPME procedures have been tested. Insample derivatization of both type of metabolites with dimethyl sulfate and sodium hydroxide was chosen as the best option because low detection limits were obtained. Moreover, laborious preconcentration steps were avoided and an important reduction of the analysis time was achieved. Experimental conditions for a high efficiency in the derivatization step were established and HSSPME parameters were optimized in order to obtain high sensitivity using the GC/MS. The optimized SPME method was compared to a SPE procedure and applied to the analysis of river water and tap water. EXPERIMENTAL SECTION Chemicals and Materials. 4-Nonylphenol (NP) technical grade was obtained from Sigma-Aldrich (Milwaukee, WI); nonyl(39) Ceglarek, U.; Efer, J.; Schreiber, A.; Zwanziger, E.; Engewald, W. Fresenius J. Anal. Chem. 1999, 365, 674-681. (40) Cuzzola, A.; Raffaelli, A.; Saba, A.; Pucci, S.; Salvadori, P. Rapid Commun. Mass Spectrom. 1999, 13, 2140-2145. (41) Cuzzola, A.; Raffaelli, A.; Saba, A.; Salvadori, P. Rapid Commun. Mass Spectrom. 2000, 14, 834-839. (42) Chee, K. K.; Wong, M. K.; Lee, H. K. J. Microcolumn Sep. 1996, 8, 131136. (43) Boyd-Boland, A.; Pawliszyn, J. Anal. Chem. 1996, 68, 1521-1525. (44) Diaz, A.; Ventura, F.; Galceran, M. T. J. Chromatogr., A 2002, 963, 159167. (45) Aranda, R.; Burk, R. C. J. Chromatogr., A 1998, 829, 401-406.

phenolmonoethoxylate (NP1EO), nonylphenoldiethoxylate (NP2EO), nonylphenoxyacetic acid (NP1EC), and nonylphenoxyethoxy acetic acid (NP2EC) were synthesized in our laboratory as described below. The derivatization reagents dimethyl sulfate (DMS), diethyl sulfate (DES), and 1-methyl-3-nitro-1-nitrosoguanidine (MNNG) were purchased from Sigma-Aldrich. Sodium hydroxide, sodium chloride, and the dechlorinating agent sodium thiosulfate were obtained from Carlo Erba (Rodano, Italy) at a high purity (g99%). The derivatization reagents, as well as NPEOsNPECs, are suspected to be carcinogenic or toxic and were handled in accordance with the most current material safety data sheets. The compounds 4n-nonylphenol (n-NP, 98%), 4n-nonyloxybenzoic acid (98%), and 4n-nonylphenolmonoethoxylate (n-NP1EO, 99%) used as internal standards were purchased from Lancaster Synthesis (Newgate, England) and Dr. Ehrenstorfer (Ausburg, Germany) respectively, whereas the surrogate for the SPE method, 4-bromophenylacetic acid, was obtained from Sigma-Aldrich. Methanol, hexane, and methylene chloride of residue analysis grade were supplied by J.T. Baker (Deventer, Holland), whereas sulfuric acid of analysis grade, anhydrous sodium sulfate, and copper(II) sulfate pentahydrate were obtained from Carlo Erba. Water from the Milli-Q water purification system (Millipore Corp., Beldford, MA) was used. SPME experiments were performed with a manual fiber holder supplied by Supelco (Bellefonte, PA). Seven commercially available fibers, (PDMS 100 and 7 µm; polyacrylate (PA) 85 µm; Carbowax-divinylbenzene (CW-DVB) 65 µm partially crosslinked and 70 µm highly cross-linked; PDMS-DVB, 65 µm; and StableFlex divinylbenzene-Carboxen-poly(dimethylsiloxane) (DVB-CAR-PDMS) 50/30 µm) were purchased from Supelco. Before use, each fiber was conditioned in a heated GC split/ splitless injection port under helium flow according to the manufacturer’s instructions. Screw-capped vials (10 and 40 mL), sealed with a Teflon-lined silicon septum and used for storing the standard solutions as well as for sample derivatization and extraction in both HS-SPME and SPE procedures, were obtained from Wheaton (Millville, NJ). The vials were cleaned by sonication with AP-13 Extran alkaline soap (Merck) for 1 h, rinsed consecutively with (i) deionized water, (ii) chromic-sulfuric acid, (iii) again with deionized water, and (iv) acetone RS grade, and baked at 50 °C overnight. Volumetric glassware was washed as described above but was air-dried. Sodium chloride and sodium hydroxide were cleaned (30 min sonication) with methylene chloride residue analysis grade and heated at 50 °C under low pressure to remove interfering organic substances. Stock standard solutions of each NPEO metabolite (1000 µg/ mL) were prepared by weight in methylene chloride. Standard mixtures were prepared weekly or daily in methanol, depending on their concentration. All solutions were stored in the dark at 4 °C until use. For the optimization of the SPME procedure, Milli-Q water samples containing NP (33 µg/L), NP1EO, NP2EO (83 µg/L each), and 165 µg/L each of NP1EC and NP2EC were prepared by adding 10 µL of a standard mixture of 100, 250, and 500 µg/ mL respectively, into 30 mL of Milli-Q water, that was kept in a 40-mL screw capped vial. Sampling Collection. Llobregat river water (northeast Spain) entering two water treatment plants located along the river course (Abrera and Sant Joan Despi (SJD)) and three tap water samples

(SJD, Llobregat, Ter) were analyzed using the HS-SPME and SPE methods. The samples were collected in 1-L amber glass bottles with PTFE-faced septa and polypropylene screw caps and stored at 4 °C. All analyses were performed within 2 days of sampling. Sodium thiosulfate (2 mL, 0.1 N) was added as a dechlorinating agent to preserve the tap water samples of free chlorine. Synthesis of Nonionic Surfactant Metabolites. NPEOsNPECs were synthesized by a new sequential method, based on well-known synthetic reactions46 and using available raw materials. NP1EC. A solution of chloracetic acid (10.4 g. 0.11 mol) in 20 mL of dimethylformamide (DMF) was added dropwise to a stirred mixture of technical nonylphenol (22 g. 0.1 mol) and NaH-60% mineral oil (4.8 g. 0.12 mol) in 30 mL of DMF. After 4 h at 65 °C, additional NaH-60% mineral oil (2.4 g. 0.06 mol) was slowly added to the mixture. The solution was stirred at 65 °C for an additional 4 h, and then it was left stirring overnight at room temperature. The resulting solution was quenched with 50% HCl (v/v). The organic layer was extracted with methylene chloride, washed with saturated NaCl solution, dried, and steam distilled. The resulting red oil was purified by chromatography using a 70-230-mesh silica gel column (20 cm × 5 cm, Fluka) and CH2Cl2-ethyl acetate, (100:1 v/v) as elution solvent. The eluate was then evaporated, and 11 g of the expected product (40% yield) was obtained. The purity (>95%) of this compound and the other metabolites that follow was established by means of 1H NMR and confirmed by GC/MS. NP1EO. A solution of NP1EC (10 g, 0.036 mol) in 50 mL of dry diethyl ether was added dropwise over a stirring mixture of 95% LiAlH4 (1.77 g, 0.044 mol) in 50 mL of dry diethyl ether at room temperature for 1 h. Lithium aluminum hydride and sodium hydride are harmful reagents and were handled in accordance with the most current material safety data sheets. The reaction was quenched by slow addition of water until no more hydrogen was formed. Then, 50% HCl (v/v) was added until pH ∼1. The organic layer was separated, dried, and evaporated in a rotary vacuum evaporator giving the expected product as a yellow oil (9 g, 90% yield). The purity of the compound was 95%. NP2EC. This compound was obtained from the previously synthesized NP1EO (6 g, 0.023 mol) using the same synthetic procedure described above for NP1EC. The resulting red oil was purified by column chromatography (silica gel, 70-230 mesh, CH2Cl2-ethyl acetate 100:1) giving the expected product (2.2 g, 30% yield). The purity of the compound was >95%. NP2EO. The synthetic procedure was the same as described above for NP1EO using the previously synthesized NP2EC (1 g, 0.003 mol). The resulting yellow oil (0.88 g, 90% yield) corresponded to the expected product and the purity was 95%. Instrumentation. The optimization of the SPME procedure was carried out with an HRGC-3000C (Konik Instruments, Barcelona, Spain) capillary gas chromatograph equipped with a FID detector, whereas quality parameters and quantitation of samples by HS-SPME and SPE were performed with a GC Fisons 8060 capillary gas chromatograph coupled to a Fisons MD 800 GC/ MS quadrupole mass spectrometer (Milan, Italy). Separations were conducted on a DB-5 MS fused-silica capillary column, 30 m × 0.25 mm i.d. × 0.25 µm (J&W Scientific, Folsom, CA), with (46) Marcomini, A.; Tortato, C.; Capri, S.; Liberatori, A. Ann. Chim. 1993, 83, 461-484.

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helium as carrier gas (70 kPa; 100 kPa N2 for FID makeup). A 0.5-m polar poly(ethylene glycol) precolumn was used. The column was held at 70 °C for 3 min, ramped at 20 °C/min to 160 °C, and finally ramped at 10 °C/min to 285 °C and held for 7 min. Splitless injection at 250 °C was used. The quadrupole mass spectrometer was operated in electron ionization (EI) positive mode (70 eV). For EI experiments, instrumental parameters were set at the following values: filament emission current of 750 µA and electron multiplier voltage of 450 V. The transfer line and the source temperature were maintained at 290 and 200 °C, respectively. The instrument was operated in SIR mode employing a dwell time of 80 ms and a delay time of 10 ms. The following ions including the base peaks of all isomers for each compound were chosen: m/z 121/135 and 149 for NP, m/z 179/193 for NP1EO, m/z 223/237 for NP2EO, m/z 207/235 and 292 for NP1EC, and m/z 117 for NP2EC. For internal standards, the monitored ions were as follows: m/z 121/234 for n-NP, m/z 231/264 for n-NP1EO, and m/z 278 for n-nonyoxylbenzoic methyl ester. For quantitation, the sum of areas for the selected ions of each methylated compound18 was monitored. Once the optimum conditions were established, different acquisition windows during each chromatographic run were applied. Masslab version 1.4 software was used for data acquisition. HS-SPME Procedure. The water sample (30 mL) was placed in a 40-mL screw-cap glass vial containing a 10 × 5 mm Tefloncoated stir bar and sodium chloride (6.5 g). After addition of 1 mL of 5 M sodium hydroxide (5 mmol), the vial was closed and 10 µL of an internal standard mixture, n-NP, n-NP1EO, and n-nonyloxybenzoic acid, of 1, 2.5, and 5 µg/mL, respectively, in methanol was added through the septum. Then, 200 µL of derivatization reagent (DMS 2.10 mmol, i.e., high excess) was introduced through the septum and the vial was clamped into a thermostated water bath at 60 °C, which was placed on a hot plate/ stirrer. A CW-DVB (70 µm) fiber was exposed to the headspace above the aqueous solution for 60 min. Magnetic stirring at 900 rpm was applied during extraction. Finally, the fiber was desorbed in the injection port of the gas chromatograph for 3 min at 250 °C. Possible carryover was prevented by keeping the fiber in the injector for an additional time (∼10 min.) with the injector in the split mode (purge on). Moreover, blanks were run periodically during the analysis to confirm the absence of contaminants. Solid-Phase Extraction Procedure. Solid-phase extraction for the determination of NPEOs-NPECs in tap water and natural water was performed following the procedure described by Ding et al.,47 with minor modifications. Briefly, acidified 500-mL samples with n-NP, n-NP1EO, and n-nonyloxybenzoic acid as internal standards (1 µg/L each) were passed through a C18 SPE cartridge (500 mg, Accubond, J&W Scientific) at a flow rate of 10 mL/min in a SPE workstation (Zymark, Hopkinton, MA). The NPEONPEC residues were eluted from the cartridge with 7 mL of a mixture of methylene chloride-methanol (9:1, v/v) acidified with acetic acid (2 mL/L). Then, the extract was evaporated to a final volume of 2 mL by a gentle stream of nitrogen. For the derivatization of NPECs, 200 µL of 96% concentrated sulfuric acid (10% of the final volume), 500 µL of hexane, and 2 µL of freshly prepared 4-bromophenylacetic acid (1 µg/mL) in methanol as derivatization surrogate standard were added to the (47) Ding, W.-H.; Chen, C.-T. J. Chromatogr., A 1999, 862, 113-120.

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final 2-mL extract. The resulting mixture was transferred to a 10mL vial sealed with a Teflon-faced septum, which was placed into a thermostated water bath at 50 °C for 1 h. After cooling to 4 °C, 5 mL of a CuSO4‚5H2O/Na2SO4 (0.2 M/0.7 M) solution was added and the mixture was shaken by hand for 2 min. An aliquot of 300 µL of the hexane extract was transferred to a 2-mL vial, and finally, 1 µL of the hexane extract was injected into the GC/MS. RESULTS AND DISCUSSION To develop a SPME method for the simultaneous quantification of short NPEOs and NPECs in water, two approaches have been tested: (i) on-fiber derivatization after extraction and (ii) in-sample derivatization previous to SPME. On-Fiber Derivatization Direct-SPME Procedure. The first strategy of derivatization48 was based on the direct extraction of the compounds from the water sample and the postderivatization of the analytes adsorbed on the fiber coating using diazomethane49,50 or BSTFA as silylation reagent.51 Different coatings were tested, and the DVB-CAR-PDMS 50/30 µm fiber appeared to be the most suitable for the extraction of the underivatized NPEOs-NPECs. However, the chromatographic profiles obtained after derivatization showed mixtures of both derivatized and underivatized compounds, making this approach insuitable. No more attempts with other reagents were assayed, and in-sample derivatization SPME was studied. In-Sample Derivatization Headspace-SPME Procedure. In a preliminary work, different derivatization reagents such as diethyl sulfate and dimethyl sulfate,52-54 methyl iodide, methyl p-toluensulfonate, or methyl trifluoromethylsulfonate were tested, to obtain the ethyl and methyl carboxylates-alcohoxylates prior to analysis by GC/MS. Direct SPME could not be used due to the extreme pH conditions of the samples that were obtained with some of these reagents and to prevent their undesirable absorption on the fiber, so HS-SPME was always employed. Among all the reagents, DMS gave the highest efficiency in the three fibers tested (PA, DVB-CAR-PDMS, CW-DVB) and therefore it was selected for the study. However, DMS confers a strong acidic character to the sample that leads to a poor derivatization yield and to an undesirable adsorption of the unreacted acidic metabolites. Moreover, damage of the chromatographic column by excess DMS on the fiber can occur. To overcome these problems, a new approach using in-sample derivatization with DMS/NaOH to form methoxy (NPEOs) and methyl ester (NPECs) derivatives and headspace sampling of the derivatized compounds has been developed. Optimization of the HS-SPME Procedure. (a) Fiber Selection. Different fiber coatings were evaluated to obtain high sensitivity and selectivity. Seven fibers were tested: PDMS 100 and 7 µm; PA 85 µm; CW-DVB 65 and 70 µm cross-linked; PDMS-DVB, 65 µm and StableFlex DVB-CAR-PDMS, 50/30 µm. (48) Li, P.; Pawliszyn, J. Anal. Chem. 1997, 69, 196-205. (49) Lee, M.-R.; Lee, R.-J.; Lin, Y.-W.; Chen, C.-M.; Hwang, B.-H. Anal. Chem. 1998, 70, 1963-1968. (50) Fales, H. M.; Jaouni, T. M.; Babashak, J. F. Anal. Chem. 1973, 45, 2302. (51) Okeyo, P. D.; Snow, N. H. J. Microcolumn Sep. 1998, 10, 551-556. (52) Neitzel, P. L.; Walther, W.; Nestler, W. Fresenius’ J. Anal. Chem. 1998, 361, 318-323. (53) Neu, H. J.; Ziemer, W.; Merz, W. Fresenius’ J. Anal. Chem. 1991, 340, 6570. (54) Sarrio´n, M. N.; Santos, F. J.; Galceran, M. T. Anal. Chem. 2000, 72, 48654873.

Figure 2. Extraction efficiency of six commercial SPME fibers. Milli-Q water containing NP (33 µg/L), NP1EO, NP2EO (83 µg/L each), and 165 µg/L NP1EC and NP2EC; DMS 200 µL; NaOH 5 M, 1 mL; sodium chloride (7.5 g); extraction time, 60 min; extraction temperature, 65 °C; and stirring rate 1200 rpm. Compound identification: (gray shaded) DVB-CAR-PDMS; (double vertical lines) CW-DVB 65 µm; (0) PA; (9) PDMS 1000 µm; (hatched) PDMS 7 µm; (checkerboard) PDMS-DVB.

A fortified water sample containing NP (33 µg/L), NP1EO, NP2EO (83 µg/L), and NP1EC and NP2EC (165 µg/L) was analyzed twice with each fiber and the following initial conditions were used: 30mL water sample (40-mL vial) with sodium chloride (7.5 g) and DMS/NaOH (ratio 1:5). Samples were heated at 65 °C, and an extraction time of 60 min was used, with the desorption temperature and desorption time of 250 °C and 5 min, respectively, for all fibers. No carryover on second desorptions were found for any of the fibers, indicating complete removal of analytes when these time and temperatures were used. The relative area responses obtained for the derivatized NPEOs and NPECs using the different fibers are shown in Figure 2. NP was the compound that gave the high preconcentration factor in all fibers except for 7-µm PDMS, because of the highest volatility of its derivatized compound. Conversely, NP2EC gave a very low values in all fibers tested. Generally, for most of the compounds, higher extraction yields were obtained for coatings with porous polymeric phases such as PDMS-DVB, CW-DVB, and the dual-coated DVB-CAR-PDMS. This may be due to the strong retention of the analytes into the pores of the polymeric phases. The CW-DVB fiber showed the highest extraction efficiency for all the compounds, and the responses were 1.1-8 times higher than those obtained with the dual-coated DVBCAR-PDMS fiber, probably because the mean micropore diameter (17 Å) of divinylbenzene is higher than Carboxen (10 Å). However, the 65-µm CW-DVB partially cross-linked fiber was too fragile at the working conditions and it could only be used for a limited number of experiments (n ∼ 10). For this reason, a 70µm CW-DVB highly cross-linked coating that keeps the same performance but has a longer lifetime was used for the remainder of the study.

Figure 3. Temperature profiles for in-sample derivatization HSSPME of NPEOs-NPECs using the CW-DVB fiber. Conditions as in Figure 2. Compound identification: ([) NP; (9) NP1EO. For (4) NP1EC, (×) NP2EO, and (O) NP2EC compounds, the relative area response is magnified by a factor of 3.

(b) Derivatization-Extraction Temperature. The effect of sample temperature on the derivatization HS-SPME was examined from 40 to 65 °C (Figure 3). An increase in the response of NP and NPEO with temperature occurred. Both the enhancement of the derivatization yield and the improvement of the mass-transfer process from the water to the headspace at high temperatures can explain the increase in the responses. Nevertheless, for NPECs, a slight decrease of the area counts at temperatures higher than 60 °C occurred. This can be attributed to a decrease of the distribution constant with temperature or the hydrolysis of methyl esters in basic media which is favored at high temperatures. As a compromise, 60 °C was chosen as the optimum temperature for the derivatization and SPME extraction of all the methyl derivatives of NPEOs and NPECs. (c) Derivatization Reagent and Sodium Hydroxide Amount. The concentration of the derivatization reagent also affects the derivatization yield of NPEOs-NPECs. To study this effect, Analytical Chemistry, Vol. 74, No. 15, August 1, 2002

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Figure 4. Optimization of derivatization reagents (DMS (mL)/NaOH 5 M (mL)). Conditions as in Figure 2, CW-DVB fiber; extraction temperature 60 °C. DMS (mL)/NaOH (mL) ratio: (gray shaded) 0.1: 0.5; (checkerboard) 0.1:1; (0) 0.2:1; (9) 0.2:2; (hatched) 0.3:1.

amounts of DMS ranging from 100 (1.0 mmol) to 300 µL (3.1 mmol) and volumes of 5 M NaOH from 0.5 (2.5 mmol) to 2 mL (10 mmol) were added to 30 mL of water spiked with the analytes at the levels indicated in the Experimental Section. The experiment was performed at 60 °C (Figure 4). In general, all volumes of DMS gave satisfactory reaction yields for most of the compounds. The amount of NaOH must be sufficient to catalyze the derivatization reaction, but it must be low enough to prevent the hydrolysis of the derivatized acidic metabolites. We observed that, for most of the compounds, mainly for the least volatile ones (NP1EC, NP2EC, NP2EO), the ratio 0.2 mL/1 mL (DMS/NaOH) gave the best results. Moreover, at these conditions, cleaner chromatograms were obtained. Thus, 1 mL of 5 M NaOH (5 mmol) and 0.2 mL of DMS (2 mmol) were the selected volumes to derivatize both NPEOs and NPECs. (d) Effect of Ionic Strength. For many organic analytes, aqueous solubility decreases with increasing ionic strength, and thus, the partitioning from the aqueous solution to the headspace is improved.35 To raise the ionic strength, an inorganic salt such as sodium sulfate54 or sodium chloride35 is often added to the aqueous matrix. In this work, the effect of ionic strength was tested with sodium chloride. An enhancement on the responses was obtained when sodium chloride concentration increased to 3.7 M (6.5 g). However, a decrease occurred at higher concentrations probably due to the high viscosity of the saline solution. So, this value was used for subsequent studies. (e) Derivatization-Extraction Time and Stirring Rate. The extraction time profiles of the methyl NPEOs-NPECs were then studied up to 75 min (Figure 5a). A total of 30 mL of Milli-Q spiked at the concentrations indicated in the Experimental Section was analyzed under the experimental conditions described in the HSSPME procedure. The adsorption time profiles for the CW-DVB fiber were established by plotting the GC-FID area response versus the extraction time for each compound in order to obtain the experimental equilibrium curve. The equilibrium time is reached when a further increase of the extraction time does not result in a significant increase in the detector response, which was established in 60 min. The effect of the stirring rate on the responses was tested between 700 and 1200 rpm. An important enhancement on the responses was obtained when the stirring rate increased from 700 to 900 rpm. At higher stirring rates (i.e., 1200 rpm), any significant increase in the area response was observed. Moreover, better precision was obtained at relatively low stirring rates than at higher ones.Thus, a stirring rate of 900 rpm was chosen for further studies. 3874 Analytical Chemistry, Vol. 74, No. 15, August 1, 2002

Figure 5. (a) Extraction time and (b) desorption temperature profiles of methyl derivatives by in-sample derivatization HS-SPME-GC-FID with CW-DVB fiber. Conditions: 30-mL spiked water sample, DMS/ NaOH (200 µL/1 mL); sodium chloride (6.5 g), extraction temperature 60 °C, and stirring rate 900 rpm. In (b) extraction time was 60 min. Compound identification: ([) NP; (9) NP1EO; (4) NP1EC; (×) NP2EO; (O) NP2EC.

(f) Optimization of Desorption Conditions. Three desorption temperatures, 210, 230, and 250 °C, within the recommended CW/DVB fiber operating range, were evaluated for a desorption time of 5 min. Results (see Figure 5b) showed differences in the GC-FID area responses between the three temperatures. Whereas the response of NPEO methyl ethers increased with temperature, a slight decrease was observed for NPEC methyl esters. Therefore, the highest temperature was chosen for further experiments. All compounds were quantitatively desorbed from the CW/DVB fiber after 3 min at 250 °C. Linearity, Precision, Sensitivity, and Matrix Effects. Quality parameters of the in-sample derivatization HS-SPME-GC/MS method were evaluated using the optimized conditions. To increase the precision, a careful selection of internal standards for HS-SPME-GC/MS was performed and n-NP, n-NP1EO, and n-nonyloxybenzoic acid,29 with structure and functional groups similar to the analytes, were used. These compounds achieved equilibrium at the same time as analytes of interest and also quantitative desorption from the fiber occurred in 3 min. By using these internal standards, the precision of the method was determined by consecutively analyzing three replicates of Milli-Q water spiked at a concentration ranging from 0.8 to 4.2 µg/L depending on the compound that was analyzed consequtively for 1 day and on three different days. Results are given in Table 1. All NPEOs-NPECs gave relative standard deviation (RSD) values for run-to-run precision lower than 8%, except for NP2EC (18%). For day-to-day precision, RSD ranged from 12 to 25%. These values were similar to those obtained by the SPE method (Table 1). The linearity of the optimized HS-SPME-GC/MS method was examined over the range 0.06-14 µg/L, depending on the compound, expressed as the initial concentration of NPEOs-NPECs in water. This range agreed with environmental levels in water currently

Table 1. Quality Parameters for In-Sample Derivatization HS-SPME-GC/MS (in Parentheses for SPE Method) precisionb,c

compound

linear range studied (µg/L)

corr coeff (r2)

LODa (µg/L)

run-to-rund

day-to-daye

NP NP1EO NP1EC NP2EO NP2EC

0.06-3.5 0.50-4.5 0.9-6.0 1.2-6.0 4.0-14

0.9952 0.9949 0.9941 0.9797 0.9737

0.02 (0.1) 0.20 (0.7) 0.30 (0.5) 0.4 (0.5) 1.5 (1.2)

8 (5) 8 (3) 7 (5) 12 (3) 18 (5)

12 (10) 15 (13) 16 (7) 22 (20) 25 (19)

a LOD limit of detection. b Precision expressed as RSDs (%). c Milli-Q water spiked at NP 0.08 µg/L; NP1EO, NP2EO 2.1 µg/L; and NP1EC, NP2EC 4.2 µg/L. d n ) 3. e n ) 3 replicates × 3 days.

Table 2. HS-SPME-GC/MS Method: Matrix Effect and Comparison with the SPE Method HS-SPME vs SPE (river water entering Abrera WTP) matrix effect (treated water)

HS-SPMEGC/MSa

SPEGC/MS a

compound

true value (µg/L)

mean (µg/L)

RSD (%) n)3

mean (µg/L)

RSD (%) n)3

mean (µg/L)

RSD (%) n)3

significance level, P-value

NP NP1EO NP1EC NP2EO NP2EC

0.83 2.1 4.2 2.1 4.2

0.76 2.1 4.0 2.0 5.1

12 19 10 12 23

1.8 0.7 2.5 1.0 11

13 17 11 19 25

1.6 0.8b 3.1 0.8b 9

6 5 16 15 7

0.065 0.153 0.072 0.141 0.141

aaIS: n-NP, n-NP1EO, and n-nonyloxybenzoic acid. bEstimated value between LOD and limit of quantification

found in the literature for these compounds. The curves were obtained by plotting the relative area of each methyl NPEONPEC to its corresponding internal standardsn-NP for NP, n-NP1EO for NP1EO, and n-nonyloxybenzoic acid for NP2EO and NPECss(A/Ais) versus the concentration of each NPEO-NPEC. The most volatile compounds showed good linearity and correlation (r2 g 0.99) although a sligthly worse linearity (r2 g 0.97) was obtained for the two less volatile compounds. Detection limits (LODs) defined as the concentration that gives an area equal to the blank plus three standard deviations were determined.. The area and the standard deviation of the blank were estimated from the calibration curve along the range studied. The limits of detection which ranged from 0.02 to 1.5 µg/L are given in Table 1 where the values obtained using the SPE procedure are also indicated. SPME leads to better LODs than SPE for the more volatile compounds especially NP, which is 5-fold lower. For the less volatile compounds, similar detection limits were obtained for both methods. LODs for the studied compounds using this procedure were slightly higher or similar than other results such as those reported for NP based on liquid-liquid extraction and derivatization of analytes with pentafluorobenzylbromide followed by GC/MS (0.001 µg/L)29 or by SPE-LC/ESI-MS30 (0.02 µg/L). Analysis of Water Samples. To examine the feasibility of the HS-SPME method, matrix effects were evaluated using a spiked free chlorine-treated water. Three replicates of a water sample were analyzed and the mean and RSD values obtained are given in Table 2. No significant differences between the mean and the true values were observed showing no interferences from other compounds potentially present in the sample. Moreover, the optimized HS-SPME method was compared to the SPE method for the simultaneous determination of NP, NPEOs, and NPECs. Triplicate samples of river water entering Abrera Water Treatment Plant were analyzed and the results are given in Table 2. To compare these results, mean values of both methods were

statistically studied using the Student’s t-test. In the case of unequal variances (F-test), the Cochran’s test was applied. The significance values (P) are given in Table 2 and showed a good agreement between the results obtained with both methods (P > 0.05). Two river water samples from the Llobregat river entering to S. Joan Despı´ (SJD) and Abrera WTPs and three tap water samples (SJD, Llobregat, Ter) were analyzed using the optimized HSSPME method. As an example, the single-ion chromatogram of methyl NPEOs and NPECs from a river water sample entering the SJD water treatment plant is displayed in Figure 6 where it can be observed that HS-SPME-GC/MS is a highly selective method for the analysis of NPEOs-NPECs. The results obtained are given in Table 3. All studied metabolites were found in Llobregat river water entering the two water treatment plants, the highest concentrations corresponded to acidic metabolites (NPECs), with NP2EC being the most abundant species. The presence of these compounds in river water is probably due to degradation of NPnEOs dumped by industries or coming from effluents of overloaded sewage treatment plants. In this context, the higher levels found in the SJD sample entering the WTP, which is situated not far from the mouth of the river, are more likely to come from dumpings from the densely industrialized area located in this part of the river. Nevertheless, in the treated water leaving the SJD water treatment plant, only nonylphenol was detected at trace levels (0.10 µg/L). The other compounds were identified at concentration levels near the LOD. This shows that the treatment that consists of prechlorination, sand filtration, groundwater dilution, ozonation, granular activated carbon filtration, and a final chlorination removes efficiently all nonylphenol metabolites. With regard to the other samples, Llobregat tap water comes from blending SJD treated water and Abrera treated water in variable amounts. The higher level of nonylphenol (0.33 µg/L) in this water sample could be explained by a less effective removal of NP in Analytical Chemistry, Vol. 74, No. 15, August 1, 2002

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Figure 6. HS-SPME-GC/MS single-ion chromatograms of derivatized compounds from river water entering SJD water treatment plant.

Table 3. Concentrations of NPEOs and NPECs in Barcelona Tap Water and River Water Samples from Different Sources by the HS-SPME-GC/MS Method SPME methoda river water (µg/L) compound

SJD

Abrera

NP NP1EO NP1EC NP2EO NP2EC

1.1 1.8 5.0 4.2 16f

1.8 0.7 2.5 1.0 11

tap water (µg/L) SJD treatedb 0.10 nd nd nd nd

Llob distribc

Ter distribd

0.33 nd nd nd nd

nde nd nd nd nd

a IS: n-NP, n-NP1EO, and n-nonyloxybenzoic acid. bSJD treated: treated water leaving SJD WTP. cLlob distrib: blended water from SJD and Abrera WTPs. dTer distrib: treated water from Ter river. e nd, not detected. fValue calculated after dilution of sample (1:2).

the Abrera WTP. Finally, no NP or the studied metabolites were found in tap water from the Ter river, thus indicating the higher raw water quality of this river or the efficient removal of these compounds in the Ter WTP. CONCLUSIONS The feasibility of HS-SPME-GC/MS for the analysis of NPEOsNPECs in water after in-sample derivatization with dimethyl sulfate/sodium hydroxide has been demonstrated. The CW-DVB 3876

Analytical Chemistry, Vol. 74, No. 15, August 1, 2002

coating was found to be the most effective for the analysis of methyl NPEOs-NPECs, especially for monoethoxylated ones. Maximum responses were obtained using 30-mL water samples salted with sodium chloride and set at an equilibration time of 60 min at 60 °C. HS-SPME in conjunction with GC/MS gave good precision, it was linear over 2 orders of magnitude, and the detection limits were at the low-microgram per liter level. This method, which avoids the use of organic solvents and requires less labor intensive sample manipulation, can be considered as an alternative to SPE and liquid-liquid extraction methods for the analysis of NP and short ethoxy metabolites in aqueous matrixes containing low concentration levels, such as drinking water. ACKNOWLEDGMENT This study was partially supported by PRISTINE Project ENV4CT97-0494 of the European Community. A.D. acknowledges a Ph.D. fellowship from Fundacio´ Agbar. SUPPORTING INFORMATION AVAILABLE Text of spectroscopic data for all synthesized compounds. This material is available free of charge via the Internet at http:// pubs.acs.org. Received for review February 26, 2002. Accepted May 13, 2002. AC020124P