Evaluation of liquid-phase microextraction conditions for determination of chlorophenols in environmental samples using gas chromatography–mass spectrometry without derivatization

Talanta 76 (2008) 154–160

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Evaluation of liquid-phase microextraction conditions for determination of chlorophenols in environmental samples using gas chromatography–mass spectrometry without derivatization Li-Wen Chung, Maw-Rong Lee ∗ Department of Chemistry, National Chung-Hsing University, Taichung 40227, Taiwan, ROC

a r t i c l e

i n f o

Article history: Received 15 November 2007 Received in revised form 15 February 2008 Accepted 18 February 2008 Available online 26 February 2008 Keywords: Without derivatization Enrichment factor Chlorophenols Liquid-phase microextraction Solid-phase microextraction Environmental samples

a b s t r a c t Determination of trace chlorophenols (CPs) in environmental samples has been evaluated using liquidphase microextraction (LPME) coupled with gas chromatography–mass spectrometry (GC–MS) without derivatization. The LPME procedure used to extract CPs from water involved 15 ␮L 1-octanol as acceptor solution in a 5.0 cm polypropylene hollow fiber with an inner diameter of 600 ␮m and a pore size of 0.2 ␮m. Under the optimal extraction conditions, enrichment factors from 117 to 220 are obtained. The obtained linear range is 1–100 ng mL−1 with r2 = 0.9967 for 2,4-dichlorophenol (2,4-DCP); 1–100 ng mL−1 with r2 = 0.9905 for 2,4,6-trichlorophenol (2,4,6-TCP); 5–500 ng mL−1 with r2 = 0.9983 for 2,3,4,6-tetrachlorophenol (2,3,4,6-TeCP), and 10–1000 ng mL−1 with r2 = 0.9929 for pentachlorophenol (PCP). The limits of detection range from 0.08 to 2 ng mL−1 , which is comparable with the reported values (12–120 ng mL−1 ). Recoveries of CPs in various matrices exceed 85% with relative standard deviations of less than 10%, except for PCP in landfill leachate. The applicability of this method was examined to determine CPs in environmental samples by analyzing landfill leachate, ground water and soil. The 2,4DCP and 2,4,6-TCP detected in the landfill leachate are 6.68 and 2.47 ng mL−1 . The 2,4,6-TCP detected in ground water is 2.08 ng mL−1 . All the studied CPs are detected in contaminated soil. The proposed method is simple, low-cost, less organic solvent used and can potentially be applied to analyze CPs in complex environmental matrices. © 2008 Elsevier B.V. All rights reserved.

1. Introduction Extensively used as preservative agents, pesticides, antiseptics and disinfectants, CPs are introduced into water either during manufacturing processes or through the degradation of phenoxyalkanoic acids. They are also formed while the chlorination of municipal drinking water [1]. Chlorophenols are carcinogenic and quite persistent [2]. The National Institutes of Health (NIH) has determined the 2,4,6-trichlorophenol (2,4,6-TCP) to be a carcinogen [3]. Therefore, CPs has been designated priority pollutants by the US Environmental Protection Agency (EPA) [4] and the European Community (EC) [5]. Exposure to large amounts of pentachlorophenol (PCP) leads to damage in the liver, kidneys, blood, lungs, nervous system, immune system and gastrointestinal tract [6–8]. Pentachlorophenol and CPs are frequently present in contaminated soils at wood treatment sites [6,9], and are very difficult to separate from various solid environmental matrices because

∗ Corresponding author. Tel.: +886 422851716; fax: +886 422862547. E-mail address: [email protected] (M.-R. Lee). 0039-9140/$ – see front matter © 2008 Elsevier B.V. All rights reserved. doi:10.1016/j.talanta.2008.02.018

they are not only firmly attached to the matrices but also soluble in non-aqueous medium. Residues of phenolic contaminants can be extracted qualitatively from solid matrices, using various extraction methods such as supercritical fluid extraction [10], Soxhlet extraction [11], shaking-assisted extraction [9], pressurized liquid extraction [12], ultrasonication-assisted extraction [13], and microwave-assisted extraction [14]. These procedures are time-consuming and require a large amount of organic solvent or expensive pretreatment equipment. Recent efforts have attempted to miniaturize liquid-liquid extraction (LLE) by markedly reducing the amount of organic solvent involved, resulting in the development of liquid-phase microextraction (LPME) methodology. Indeed, LPME reduces 99% less organic solvent than traditional LLE [15]. Since LPME requires micro-volume of solvent, applications of this method have attracted considerable attention and are feasible to environment [16–24]. In addition to minimizing organic solvent, high enrichment is also one factor to validate LPME while extraction. In other way, high enrichment is needed to enhance the sensitivity for CPs when analyzing environmental samples. Recently, determination of CPs by 3-phase LPME coupled with HPLC [17] and aqueous samples were

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extracted by LPME with high enrichment factors. This provides an opportunity for quantification of the LPME method by implementing the kinetic approach. Otherwise, a derivatization step enhances sensitivity particularly when GC was used for the determination of CPs. The derivatization leads to sharper peaks and hence to better separation and higher sensitivity for the CPs. Different derivatization systems have been used including water-bath derivatization [25,26], in situ derivatization [7,27], injection port-derivatization [28] and microwave-assisted derivatization [29]. Although derivatization is one of the most common procedures to enhance the sensitivity of CPs, it unnecessarily contaminates the GC–MS system, shortens column life and creates a need for additional maintenance. In order to avoid these problems, the method without derivatization is used as modified conditions to get high enrichment factor for trace CPs determination. Therefore, this study explores the potential of procedure without derivatization based on LPME to achieve high enrichment factor. Since LPME is a process dependent on equilibrium [20,30], the objective of this work is to reach high distribution equilibrium in the extraction system. Herein, the optimal conditions were examined systematically to evaluate linearity, limits of detection, limits of quantification and precision. Furthermore, the validity of this method was also applied to determine CPs in contaminated environmental samples and assessed with our previous solid-phase microextraction (SPME) method [31]. 2. Experimental 2.1. Chemicals, reagents and materials 2,4-Dichlorophenol (2,4-DCP, 99%), 2,4,6-TCP (97%), 2,3,4,6tetrachlorophenol (2,3,4,6-TeCP, 80%) were obtained from TCI (Tokyo, Japan). 2,3,4,6-TeCP was further purified by recrystallization from diethyl ether and a final purity of over 99% obtained [31]. 2,4,6-Tribromophenol (99%) as a surrogate standard was obtained from TCI and used without further purification. Pentachlorophenol (neat) was purchased from Supelco (Bellefonte, PA, USA). A stock solution of a mixture of CPs at 100 mg L−1 for 2,4-DCP, 100 mg L−1 for 2,4,6-TCP, 500 mg L−1 for 2,3,4,6-TeCP and 1000 mg L−1 for PCP was prepared by dissolving the solid bulk CPs in isopropanol (Merck, Darmstadt, Germany) and storing it at 4 ◦ C. The stock solution was further diluted to yield the appropriate working solutions with water. 1-Octanol (99.5%) and NaCl (99.8%) were purchased from Riedel¨ (Seelze, Germany). KCl (99%) and hydrochloric acid (HCl, deHaen 37.5%) were ordered from Fisher (Fair Lawn, NJ, USA). Citric acid ¨ NaOH (98.9%) was from (99.5%) was bought from Riedel-deHaen. TEDIA (Fairfield, Ohio, USA). The laboratory purified water (18 M) was obtained using a SG-Ultra Clear water purification system (SG, ˝ Barsbuttel, Germany). The pH 1 buffer solution was prepared by using 0.1 M KCl and adjusted to pH 1 with 0.2 M HCl. The citrate solution was prepared by adding 200 mL of 1 M NaOH solution to dissolve 21.014 g of citric acid, and diluting to 1000 mL with pure water. To obtain buffer solutions with pH values between 2 and 4, suitable volumes of 0.1 M HCl were added to citrate solution. To obtain buffer solutions with pH values 5 and 6, suitable volumes of 0.1 M NaOH were added to citrate solution. 2.2. GC–MS analysis Chromatographic analysis was performed using a HewlettPackard (HP) MS Engine mass spectrometer (Palo Alto, CA, USA) with an HP 5890 Series II GC through an injector in splitless mode. Separations were carried out using a 30 m × 0.25 mm fused cap-

155

Table 1 Analytical conditions of chlorophenols determined by GC–MS Compound

Retention time (min)

Quantification ion/confirmation ion (m/z)

Mr

2,4-DCP 2,4,6-TCP 2,3,4,6-TeCP PCP

2.38 2.72 3.50 4.59

162/164 (3:2) 196/198 (1:1) 232/230 (4:3) 266/264/268 (15:9:10)

162 196 230 264

illary column DB-5MS (J&W Scientific, Folsom, CA, USA) with a stationary phase thickness of 0.25 ␮m. Helium (99.999%) was used as the carrier gas at a constant flow of 1 mL min−1 . In LPME, the temperature of the injector port was maintained at 250 ◦ C. The oven was initially set to 130 ◦ C, programmed to 190 ◦ C at a rate of 30 ◦ C min−1 , and then to 230 ◦ C at rate of 10 ◦ C min−1 . The total analysis time for a single run took 6 min. The GC–MS temperature of the transfer line was maintained at 250 ◦ C. In SPME, the temperature of the injector port was maintained at 290 ◦ C. The oven was initially set to 60 ◦ C, programmed to 190 ◦ C at a rate of 30 ◦ C min−1 , and then to 310 ◦ C at a rate of 10 ◦ C min−1 . Here, the total analysis time for a single run was 16 min. The GC–MS temperature of the transfer line was maintained at 310 ◦ C [31]. The ion source and quadrupole temperatures were set to 250 ◦ C and 100 ◦ C, respectively. Mass spectra were obtained using the electron impact (EI) mode. The full scan mode with a mass range of m/z 40–350 was adopted to confirm the CPs and the selected ion monitoring (SIM) mode was applied to quantify the CPs. The most abundant ion was used as the quantification ion and the specific ion was used as the confirmation ion, as indicated in Table 1. 2.3. Liquid-phase microextraction The experimental set-up for extraction was described as reported by Pedersen-Bjergaard and Rasmussen [30]. The Accurel® Q3/2 polypropylene hollow fiber (600 ␮m inner diameter, 200 ␮m wall thickness, 0.2 ␮m pore size) was purchased from Membrana (Wuppertal, Germany). Before extraction, the hollow fiber was ultrasonically cleaned in acetone for 30 min to remove contaminant. After it had been dried, the hollow fiber was cut by hand into 5.0 cm lengths. Briefly, a fresh hollow fiber was immersed in 1-octanol for approximately 5 min to impregnate the pores of the fiber with the solvent and then known amounts of 1-octanol (15 ␮L) as acceptor solution was injected carefully into the hollow fiber by syringe. The hollow fiber filled with 1-octanol was then immersed for extraction into 30 mL of donor solution, which was prepared by mixing 15 mL of pH buffer solution with 15 mL of sample solution (contained 5 ␮g mL−1 surrogate standard). The donor solution was continuously stirred using a magnetic stirrer to facilitate the mass transfer process and reduce the time required to reach equilibrium (80 min). After the extraction time, the acceptor solution in the hollow fiber was withdrawn by the syringe and collected in a 0.7 mL conical vial. A 1 ␮L of extraction solution was taken to analyze. 2.4. Solid-phase microextraction The SPME condition was according with our previous work [31]. In briefly, the experiment was conducted using a commercially available polyacrylate fiber with a film thickness of 85 ␮m, housed in its manual holder (Supelco, Bellefonte, PA, USA). The new polyacrylate fiber was conditioned using the injector port of the GC at 300 ◦ C for 2 h. During extraction, 12.5 mL of the sample was added to 12.5 mL of pH 1 buffer solution saturated with KCl for adsorption 40 min at a stirring rate of 1000 rpm. Then, the extract was

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desorbed from the fiber into a GC injector at 290 ◦ C at the maximum length (4.5 cm) of the syringe carriage for 2 min. 2.5. Validation of method Blank samples were spiked with 2,4-DCP at 1, 25, 50, 75 and 100 ng mL−1 , 2,4,6-TCP at 1, 25, 50, 75 and 100 ng mL−1 , 2,3,4,6-TeCP at 5, 125, 250, 375 and 500 ng mL−1 and PCP at 10, 250, 500, 750 and 1000 ng mL−1 , then analyzed by the optimum LPME procedure to generate calibration curves. These curves were obtained by plotting the peak area ratio (analyte to 5 ␮g mL−1 surrogate standard) as a function of the concentration. The limit of detection (LOD) was defined as the concentration in the sample that resulted in a peak with an S/N (signal-to-noise ratio) of three [32]. The limit of quantification (LOQ) was defined as the concentration in the sample that yielded a peak with an S/N of ten [32]. The precision of the assay was evaluated by analyzing the CPs in quality control (QC) samples on the same day. Quality control samples were prepared at three different concentrations of low, medium and high. The low QC concentration sample: 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP were at 1, 1, 5 and 10 ng mL−1 , respectively. The medium QC concentration sample: 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP were at 50, 50, 250 and 500 ng mL−1 , respectively. The high QC concentration sample: 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP were at 100, 100, 500 and 1000 ng mL−1 , respectively. 2.6. Sample preparation A soil sample was obtained from the CPs-contaminated soil at an abandoned chemical manufacturing site in Tainan (southern of Taiwan). After the soil was shaken through two mesh screens with mesh sizes of 1.981 mm and 2.000 mm, it was collected and stored at 4 ◦ C in a refrigerator. The sample solution was prepared by soaking 30 mg of soil in 15 mL aqueous solution that contained 5 ␮g mL−1 surrogate standard. Landfill leachate was obtained from a landfill site in Taichung (middle of Taiwan). Ground water was collected from a ground water tap, which was allowed to flow for 10 min. The landfill leachate sample solution was prepared by 15 mL landfill leachate contained 5 ␮g mL−1 surrogate standard. The ground water sample solution was prepared similar to the landfill leachate samples. 3. Results and discussion

Fig. 1. Effect of pH on peak area of chlorophenols obtained by LPME–GC–MS. Experiment conditions: 100 ng mL−1 of 2,4-DCP (), 100 ng mL−1 of 2,4,6-TCP (), 500 ng mL−1 of 2,3,4,6-TeCP () and 1000 ng mL−1 of PCP (); without the addition of NaCl; stirring rate: 550 rpm; extraction time: 30 min.

that the extraction efficiency of the four CPs decreases as the NaCl concentration increases. This phenomenon is explained by the fact that the electrostatic interactions between analytes and the salt ions in the donor solution are stronger than the salting out effect. Hence, no NaCl was added in the following studies. 3.3. Effect of stirring rate on extraction efficiency Extraction kinetics can be accelerated by agitating samples. Here donor solutions were examined at various stirring rates for 30 min to evaluate the effect of the stirring rates. As shown in Fig. 3, the higher the stirring rate, the better the extraction efficiency for all the studied CPs. This result is consistent with the film theory of convective diffusion [34] that attributes the high extraction efficiency to the continuous exposure of fresh donor solution. Since the extracted amount reached the highest value at 1100 rpm, the same stirring rate was used for ensuing studies. 3.4. Effect of extraction time on extraction efficiency Liquid-phase microextraction is an equilibrium process between donor solution and acceptor solution [20,30]. Based on

3.1. Effect of pH on extraction efficiency The pH value relates to the equilibrium between the ionized form and the neutral form. Chlorophenols are extracted mainly as neutral molecules in LPME. The results in Fig. 1 indicate that the extraction efficiency increases as the pH value decrease. At low pH, the equilibrium of CPs favors neutral form, which is more soluble in the acceptor solution and has a lower affinity toward donor solution. According to these results, the pH value was adjusted to one in subsequent experiments. 3.2. Effect of salt on extraction efficiency According to Zhao et al. [33], the effect of salt on extraction can be separated into two simultaneous processes—salting out and electrostatic interactions. The salting out effect increases the amount of extraction at high salt concentration, depending on the solubility of the analytes. The electrostatic interaction method decreases the amount of extraction at high salt concentration, involving attractive forces between charged particles that are dispersed in electrolytes and the analytes. The results in Fig. 2 reveal

Fig. 2. Effect of salt addition on peak area of chlorophenols obtained by LPME–GC–MS. Experiment condition: 100 ng mL−1 of 2,4-DCP (), 100 ng mL−1 of 2,4,6-TCP (), 500 ng mL−1 of 2,3,4,6-TeCP () and 1000 ng mL−1 of PCP (); pH 1 buffer; stirring rate: 550 rpm; extraction time: 30 min.

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Fig. 3. Effect of stirring rate on peak area of chlorophenols produced by LPME–GC–MS. Experiment condition: 100 ng mL−1 of 2,4-DCP, 100 ng mL−1 of 2,4,6TCP, 500 ng mL−1 of 2,3,4,6-TeCP and 1000 ng mL−1 of PCP; pH 1 buffer; without the addition of NaCl; extraction time: 30 min.

partition coefficient of CPs, this characteristic is particularly beneficial in quantification of the LPME method. It enables substantial portion of the CPs extracted from donor solution to acceptor solution. For this reason equilibrium time was studied herein was 10–180 min. For all studied CPs, the amount extracted increases markedly with the extraction time from 10 to 80 min, as shown in Fig. 4. At 80 min, the amount of extracted CPs reached equilibrium. Therefore, 80 min was used as the optimal extraction time in further analysis. Compared the extraction time with previous literature, which was 30 min [28] and 60 min [17], this developed method is comparatively long may be due to the slow mass transfer for analytes into organic solvent [35]. 3.5. Enrichment factor The enrichment factor can be defined as the ratio of the equilibrium concentration of analytes in the organic phase to the original concentration of analytes in the aqueous phase [36]. Table 2 shows that the enrichment factor in this procedure varies from 117 to 220, depending on the CPs studied. The high enrichment factor is obtained through the optimum conditions, which are mixing 15 mL of pH 1 buffer solution with 15 mL of sample solution with a sur-

Fig. 4. Effect of extraction time on peak area of chlorophenols produced by LPME–GC–MS. Experiment condition: 100 ng mL−1 for 2,4-DCP (), 100 ng mL−1 of 2,4,6-TCP (), 500 ng mL−1 of 2,3,4,6-TeCP () and 1000 ng mL−1 of PCP (); pH 1 buffer; without the addition of NaCl; stirring rate: 1100 rpm.

157

rogate standard concentration of 5 ␮g mL−1 at 1100 rpm for 80 min extraction. High enrichment factor is evidence for high partition coefficients of the anlytes and outstanding extraction efficiency in LPME. With the high enrichment factor, this LPME method is capable of reaching low enough detection ranging from 0.08 to 2 ng mL−1 and quantification limits between 0.3 and 7 ng mL−1 . Compared with other extraction technique, the LOD was 12 ng mL−1 for 2,4-DCP by ultrasound-assisted headspace LPME [16], 0.5 ␮g L−1 for 2,4-DCP and 1.0 ␮g L−1 for 2,4,6-TCP by hollow fiber supported ionic liquid membrane extraction [17], and 53 ng g−1 for 2,4-DCP, 12 ng g−1 for 2,4,6-TCP, 39 ng g−1 for 2,3,4,6-TeCP and 120 ng g−1 for PCP by focused microwave-assisted micellar extraction combined with SPME [37]. It could be seen that the LODs of the proposed method are fairly better than that obtained by previous works. This result is in agreement with that obtained by Wen et al. [38], which the advantages of high enrichment is capable of achieving lower detection. 3.6. Performance of method The linearity, LOD, LOQ and precision were investigated in water under the aforementioned optimal conditions of LPME coupled to GC–MS procedure, as shown in Tables 2 and 3. The correlation coefficient of each studied analyte is above 0.99 in the range of interest, but correlation coefficient gives insufficient information about the linearity of the curve. A linearity test is performed to give sufficient information about the linearity of curve by comparing the linear calibration function with the nonlinear calibration function. According to DIN 38402 section 51, a test value (TV) was calculated from the residual standard deviation [39]. The TV value was smaller than the F-test value as shown in Table 2; therefore the calibration function is linear. The detection response factor (RF) was used to calculate the coefficient of variation (CV). According to Telliard et al. [40], a RF is acceptable if CV < 20%. The CV values calculated are presented in Table 2, where all CVs calculated were less than 10%. Based on the linearity test and considering the small standard error of slope, the analysis of the breakthrough samples was performed using a single calibration curve over the whole concentration range. After generating calibration curves as depicted in Table 2, the linear regression analysis for CPs is observed on the slopes between 0.0035 and 0.0055, intercepts between 0.0483 and 0.1076. With the advantages of high enrichment factor, the obtained LOD and LOQ are in the range of 0.08–2 ng mL−1 and 0.3–7 ng mL−1 , respectively, competitive with the reported LOD in literature, which were 12–120 ng g−1 [37]. Precision in terms of repeatability and reproducibility is expressed as the relative standard deviation (R.S.D.) [41]. Repeatability is the degree of agreement among individual test results when the procedure is applied repeatedly. It was measured from six replicate analyses of the QC samples. As seen in Table 3, the R.S.D.s ranged between 1.15 and 8.87% for repeatability. Even in the low QC concentration, the R.S.D.s are lower than 10% for repeatability. These results for repeatability show the suitability of the LPME–GC–MS technique in the analysis of trace CPs. In the case of reproducibility, six replicates were analyzed for 3 consecutive days at QC samples. The R.S.D.s ranged from 2.92 to 11.63% for reproducibility, as indicated in Table 3. The R.S.D.s of reproducibility are within ±15% for all measurements, it shows the good precision of the method [41]. 3.7. Real sample In order to overcome the difficult extraction from soil, LPME was applied to determine the CPs from the CPs-contaminated soil. This

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Table 2 Calibration curve results, limits of detection (LOD), limits of quantification (LOQ) and enrichment factor of chlorophenols in water produced by LPME–GC–MS Compound

Correlation coefficient

TV

CV (%)

Linear range (ng mL−1 )

Linear equation

LOD (ng mL−1 )

LOQ (ng mL−1 )

Enrichment factor

2,4-DCP 2,4,6-TCP 2,3,4,6-TeCP PCP

0.9967 0.9905 0.9983 0.9929

0.30 0.05 0.28 0.02

6.29 9.87 3.77 8.91

1–100 1–100 5–500 10–1000

Y = 0.0046X + 0.1018 Y = 0.0055X + 0.1076 Y = 0.0040X + 0.0956 Y = 0.0035X + 0.0483

0.10 0.08 1.31 2.01

0.33 0.26 4.37 6.70

176 117 155 220

F-test (0.99, 1, 2) = 98.5.

Table 3 Repeatability and reproducibility for chlorophenols in water spiked at three QC concentrations in water produced by LPME–GC–MS (n = 6) Compound

2,4-DCP 2,4,6-TCP 2,3,4,6-TeCP PCP

Repeatability (R.S.D., %)

Reproducibility (R.S.D., %)

Low QC concentration

Medium QC concentration

High QC concentration

Low QC concentration

Medium QC concentration

High QC concentration

3.28 8.87 7.21 4.05

1.85 1.61 1.51 1.37

1.15 1.34 1.58 1.70

10.39 9.14 11.63 9.26

6.71 3.26 3.06 6.32

2.92 3.21 3.52 6.43

Low QC concentration was spiked 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP at 1, 1, 5 and 10 ng mL−1 , respectively. Medium QC concentration was spiked 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP at 50, 50, 250 and 500 ng mL−1 , respectively. High QC concentration was spiked 2,4-DCP, 2,4,6-TCP, 2,3,4,6-TeCP and PCP at 100, 100, 500 and 1000 ng mL−1 , respectively.

method was also utilized to extract CPs from a complex matrix, such as landfill leachate and ground water. Table 4 demonstrates the concentration of CPs determined from the environmental samples by LPME. Two analytes, 2,4-DCP and 2,4,6-TCP were detected in the landfill leachate and the concentrations were 6.68 and 2.47 ng mL−1 , respectively. In ground water, only 2.08 ng mL−1 of 2,4,6-TCP was detected. In the case of soil samples, all the studied CPs were detected, as seen in Table 4. To assess quantification analysis, the measured result of LPME is compared with the result obtained by SPME for the environment sample. The SPME condition was according with our previous work

[31]. The determined concentrations show no variation, as depicted in Table 4. Furthermore, fewer significant interference peaks in chromatogram (Fig. 5b and c) were obtained than those obtained by using SPME (Fig. 5e and f). Base on the quantification and chromatogram of LPME, it clearly reveals that the LPME–GC–MS technique exactly quantifies CPs and effectively reduces the matrix interference in the environmental samples. The components of the environmental sample are very complex and the matrix effect reduces the recovery. The environmental samples were spiked with all CPs at medium QC concentration to real samples in order to evaluate the recovery. The recovery is calculated

Fig. 5. Mass ion chromatograms (a) 25 ng mL−1 of 2,4-DCP, 25 ng mL−1 of 2,4,6-TCP, 50 ng mL−1 of 2,3,4,6-TeCP and 250 ng mL−1 of PCP in water solution produced by LPME–GC–MS; (b) landfill leachate produced by LPME–GC–MS; (c) contaminated soil produce by LPME–GC–MS; (d) 25 ng mL−1 of 2,4-DCP, 25 ng mL−1 of 2,4,6-TCP, 50 ng mL−1 of 2,3,4,6-TeCP and 250 ng mL−1 of PCP in water solution produced by SPME–GC–MS (e) landfill leachate produced by SPME–GC–MS; (f) contaminated soil produced by SPME–GC–MS.

5.13 1.16 4.54 3.18 93.02 90.52 106.47 99.86 7.72 22.93 32.16 46.90 3.45 9.50 4.30 4.28 91.39 94.75 109.53 101.31

4. Conclusions

6.68 2.47 ND ND

97.21 87.38 89.45 65.85

10.30 3.45 1.24 2.84

ND 2.08 ND ND

The proposed method is highly successful for determining trace amounts of CPs in environmental samples using LPME coupled with GC–MS without derivatization. Factors that influence the extraction efficiency have been investigated. High enrichment factor is sufficient to establish the outstanding extraction efficiency. The LODs of CPs are 0.08–2 ng mL−1 , revealing the high sensitivity of this method and comparable with the reported value (12–120 ng mL−1 ). Under optimal conditions, recoveries of CPs in environmental samples exceed 85% with R.S.D.s of less than 10%, except for PCP in landfill leachate. Liquid-phase microxtraction coupled with GC–MS is favored over SPME and involves no carry-over problem or fewer interference peaks. Based on the simplicity and sensitivity, this method can be recommended to determine CPs in environmental samples.

4.71 6.45 7.83 8.66 89.43 93.26 101.88 86.86 8.19 17.08 38.86 48.79

159

from the ratio of the peak area obtained in environmental sample to that in pure water. In Table 4, the recoveries of SPME were between 7.44 and 58.18% in landfill leachate, 72.74 and 94.22% in ground water, 86.86 and 101.88% in soil. For LPME, the recoveries were between 65.85 and 97.21% in landfill leachate, 91.39 and 109.31% in ground water, 90.52 and 106.47% in soil. In contrast with Peng et al. [17], the recovery was 90.1% for 2,4-DCP and 88.7% for 2,4,6-TCP in ground water. In present study, the recovery in ground water was 91.39% for 2,4-DCP and 94.75% for 2,4,6-TCP. The R.S.D. of SPME was below 11.58% and the R.S.D. of LPME was below 10.30%. Since the LPME recoveries are higher than SPME with smaller R.S.D. value for all environmental samples, it clearly demonstrates that LPME effectively prevent matrix effect from complex environmental matrices.

Acknowledgments The authors would like to thank the National Science Council of the Republic of China, Taiwan, for financially supporting this research under Contract No. NSC 94-2113-M-005-001. K. David Tau is appreciated for his editorial assistance.

7.44 6.84 9.74 11.58

ND 2.39 ND ND

80.66 86.83 94.22 72.74

5.56 5.78 8.17 9.92

References

a

ND: Not detected.

58.18 16.25 8.81 7.44 6.35 2.72 NDa ND 2,4,-DCP 2,4,6-TCP 2,3,4,6-TeCP PCP

Soil

Conc. (ng mL−1 )

Ground water

Conc. (ng mL−1 ) Recovery R.S.D. (%) (%)

Landfill leachate

Conc. (ng mL−1 ) Recovery R.S.D. (%) (%) Conc. (ng mL−1 ) Recovery R.S.D. (%) (%) Conc. (ng mL−1 ) Recovery R.S.D. (%) (%) Conc. (ng mL−1 )

LPME

Soil Ground water Landfill leachate

SPME Compound

Table 4 Comparison of LPME with SPME on environmental samples (n = 3)

Recovery R.S.D. (%) (%)

Recovery R.S.D. (%) (%)

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