Anal Bioanal Chem (2004) 378 : 1854–1860 DOI 10.1007/s00216-004-2510-8

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O R I G I N A L PA P E R

Hiroaki Otaka · Hisao Shimono · Shunji Hashimoto

Optimization of solid-phase extraction procedure for determination of polychlorinated dibenzo-p -dioxins, polychlorinated dibenzofurans, and coplanar polychlorinated biphenyls in humic acid containing water Received: 31 October 2003 / Revised: 5 January 2004 / Accepted: 14 January 2004 / Published online: 12 February 2004 © Springer-Verlag 2004

Abstract A solid-phase extraction (SPE) method was optimized for accurate determination of polychlorinated dibenzo-p-dioxins (PCDDs), polychlorinated dibenzofurans (PCDFs), and coplanar polychlorinated biphenyls (CoPCBs) in humic acid containing surface water. Recovery experiments using humic materials revealed that humic acids permit dioxins to pass through an octadecylsilica (C18) extraction disk by associating with them under weakly alkaline conditions. Acidification of the sample before percolation improved this otherwise insufficient recovery. The analysis of surface water acidified to pH 2 gave better recovery with surrogate standards and lower quantitative values for higher-chlorinated homologues than the sample at pH 9. In all samples, the native octachlorodibenzo-p-dioxin (OCDD) peak abundance showed no difference between at pH 2 and at pH 9, indicating overestimation of the quantitative value of the homologue at pH 9. Acidification of a humic acid containing water sample can avoid overestimation of higher-chlorinated congeners caused by insufficient recovery of their corresponding surrogates. Keywords Polychlorinated dibenzo-p-dioxins · Polychlorinated dibenzofurans · Coplanar polychlorinated biphenyls · Surface water · Solid-phase extraction · Humic substances

H. Otaka (✉) National Environmental Research and Training Institute, Ministry of the Environment, 3–3 Namiki, Tokorozawa-shi, 359–0042, Saitama, Japan e-mail: [email protected] H. Shimono Japan Quality Assurance Organization, 14–12 Dezuminato, Chuo-ku, Chiba-shi, 260–0023 Chiba, Japan S. Hashimoto National Institute for Environmental Studies, 16–9 Onogawa, Tsukuba-shi, 305–8506 Ibaraki, Japan

Introduction Polychlorinated dibenzo-p-dioxins (PCDDs), polychlorinated dibenzofurans (PCDFs), and polychlorinated biphenyls (PCBs) are chemically stable and present in almost all kinds of environmental samples such as soil, fly ash, sewage sludge, water, etc. [1]. Of these compounds, PCDD/F homologues with chlorine atoms in the 2,3,7,8-positions and dioxin-like coplanar PCBs (CoPCBs) are regarded as the most toxic. Precise estimation of their concentrations in environmental samples is required for assessment of their risk to human health. Analysis of dioxins in water is a challenging task, since the concentrations of these compounds are extremely low due to their low solubility in this medium [2]. For this reason, larger-volume water samples and higher-sensitivity analytical instruments are needed to reach the lower detection limits of the target analytes. Most conventional methods, such as the Japanese JIS K0312 [3] and the US EPA method 1613 [4], permit either liquid–liquid extraction (LLE) or solid-phase extraction (SPE) using an octadecylsilica (C18) membrane as an extraction step for determination of dioxins in water. The SPE method consists of prefiltration of a sample via a filter, and percolation of the filtrate with a C18 membrane disk. For extraction from high volumes of water, SPE is faster and requires lower volumes of organic solvents than LLE. The SPE technique has several advantages as mentioned above, but also suffers from several drawbacks including clogging and overloading. Of these problems, the insufficient recovery of surrogate standards added to samples before processing is of particular concern. When determining dioxins in surface water by SPE, most samples give poor rates of recovery (below 70%) of the surrogates of higher-chlorinated PCDD/Fs, suggesting the existence of a factor responsible for loss of the target analytes in environmental water. This study focuses on humic substances, which are major components of dissolved organic carbon (DOC) in environmental water [5]. Humic acids (HAs) have common

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specific characteristics, that is, they have domains with hydrophobic character where nonpolar pollutants can be trapped, while other parts present carboxylic and hydroxyl groups which, depending on the pH, are more or less protonated. Studies have been made on the interaction of the hydrophobic domains with nonionic compounds such as PCDD/Fs [6], polycyclic aromatic hydrocarbons (PAHs) [7, 8, 9], PCBs [10], and several kinds of pesticides [10, 11]. It has been also reported that owing to its polyionic properties, HA affects the retention of several pesticides or PAHs in reversed-phase cartridges [9, 12]. These reports suggest that HA in surface water may be responsible for the observed insufficient recovery of dioxin surrogates. We anticipated that the chance to extract the maximum amount of HA molecules bearing nonpolar compounds with a hydrophobic C18 solid phase may be when the HA molecules are weakly polar or, in other words, when they have a maximum number of protonated groups (i.e., at low pH). We have tried to verify this point in the case of the extraction of PCDD/Fs and CoPCBs with a C18 solid phase. The objectives of this work were (i) to investigate the effects of humic substances on the recovery of PCDD/Fs and CoPCB homologues when using solid-phase extraction, (ii) to optimize an SPE procedure with a view of obtaining sufficient recovery of surrogates for determination of dioxins in surface water. Such accurate analytical procedures should be indispensable for the reasonable estimation of the environmental fate or toxicological impact of dioxins on aquatic organisms. Sufficient recovery of surrogates is one criterion for achieving accurate determination of dioxins. As will be seen, solving the abovementioned problem is of fundamental importance in obtaining accurate quantitative values of dioxins in humic acid containing water.

Experimental Apparatus A continuous sample absorption system purchased from GL Sciences (Tokyo, Japan) was used for sample prefiltration with a glass fiber filter, and percolation was performed with an SPE disk. Pressurized liquid extraction (PLE) with acetone was performed using ASE-300 (Dionex, CA, USA) for extraction of the target analytes from a glass fiber filter and a C18 membrane. The extraction conditions were as follows: cell volume 66 mL, cell temperature 150°C, cell pressure 10 MPa (1,500 psi), static time 7 min, solvent flush 66 mL (100% of cell volume), 100-s purge with N2 gas, and 2 cycles. Analysis of PCDD/Fs and CoPCBs was performed by high-resolution gas chromatography–high-resolution mass spectrometry (HRGC-HRMS) using an HP-6890 plus (Agilent, CA, USA) gas chromatograph coupled to a JMS-700 mass spectrometer (JEOL, Tokyo, Japan). The determination of the analytes was performed in BPX-DXN (SGE, TX, USA) and RH-12 ms (Inventx, CA, USA) capillary columns. Determination of DOC in water samples was performed using TOC-5000A (Shimadzu, Kyoto, Japan).

Samples, materials, and reagents Four surface water samples used in the quantitative experiments were collected from the stagnant spot of a river (samples 1–3) or a pond (sample 4) in the Saitama prefecture, Japan. The Empore C18-FF disk (90-mm i.d., 0.5-mm thick) was purchased from 3M (MN, USA). The membrane was prewashed by Soxhlet extraction with toluene for 16 h prior to percolation. The GB-100R glass fiber filter (0.6-µm particle retention) was obtained from Toyo Roshi (Tokyo, Japan). All analytical-grade solvents and adsorbents of dioxins were purchased from either Wako Pure Chemical Industries (Osaka, Japan) or Kanto Chemicals (Tokyo, Japan). Analytical-grade hydrochloric acid, sodium hydroxide, and potassium dihydrogen phosphate were purchased from Kanto Chemicals. PCDDs/PCDFs and CoPCBs standards including 13C12-labeled compounds were obtained from Wellington Laboratories (Ontario, Canada) or Cambridge Isotope Laboratories Inc. (MA, USA). In the following text and all tables, the number of chlorines present from 4 to 8 are given as tetra (Te), penta (Pe), hexa (Hx), hepta (Hp), and octa (O), for example, 2,3,7,8-TeCDD represents tetrachlorodibenzo-p-dioxin with chlorines in positions 2, 3, 7, and 8. All CoPCBs were represented by the International Union of Pure and Applied Chemistry (IUPAC) numbers. A surrogates solution (SS) was prepared in acetone. This mixture contains 13C-labeled 2,3,7,8TeCDD (TeCDD1), 1,2,3,7,8-PeCDD (PeCDD1), 1,2,3,4,7,8-HxCDD (HxCDD1), 1,2,3,6,7,8-HxCDD (HxCDD2), 1,2,3,7,8,9-HxCDD (HxCDD3), 1,2,3,4,6,7,8-HpCDD (HpCDD1), OCDD, 2,3,7,8TeCDF (TeCDF1), 1,2,3,7,8-PeCDF (PeCDF1), 2,3,4,7,8-PeCDF (PeCDF2), 1,2,3,4,7,8-HxCDF (HxCDF1), 1,2,3,6,7,8-HxCDF (HxCDF2), 1,2,3,7,8,9-HxCDF(HxCDF3), 2,3,4,6,7,8-HxCDF (HxCDF4), 1,2,3,4,6,7,8-HpCDF (HpCDF1), 1,2,3,4,7,8,9-HpCDF (HpCDF2), OCDF, 3,3′,4,4′-TeCB (#77, TeCB1), 3,4,4′,5-TeCB (#81, TeCB2), 2,3,3′,4,4′-PeCB (#105, PeCB1), 2,3,4,4′,5-PeCB (#114, PeCB2), 2,3′,4,4′,5-PeCB (#118, PeCB3), 2′,3,4,4′,5-PeCB (#123, PeCB4), 3,3′,4,4′,5-PeCB (#126, PeCB5), 2,3,3′,4,4′,5HxCB (#156, HxCB1), 2,3,3′,4,4′,5′-HxCB (#157, HxCB2), 2,3′,4,4′,5,5′-HxCB (#167, HxCB3), 3,3′,4,4′,5,5′-HxCB (#169, HxCB4), and 2,3,3′,4,4′,5,5′-HpCB (#189, HpCB1) at the concentration of 1.0 pg µL–1 each, with the exception of 13C12-OCDD/F which was at 2.0 pg µL–1. A recovery standards solution (RSS) containing 13C-labeled 1,2,7,8-TeCDF, 1,2,3,4,7-PeCDD, 1,2,3,4,6,9HxCDF, 1,2,3,4,6,8,9-HpCDF, and 2,3′,4′,5-TeCB (#70) was prepared in nonane at the concentration of 10 pg µL–1 each. An extraction standard was also used to monitor the extraction efficiency and the loss of target analytes during cleanup of the extract. This standard solution (ESS), containing 13C12-1,2,3,4,6,7-HxCDD, was prepared at the concentration of 10 pg µL–1 in nonane. A well-characterized HA standard and a fulvic acid (FA) standard, both derived from forest soil, were provided by the Japanese Humic Substances Society (JHSS). The commercial HA product used in this study was purchased from Wako. A stock solution of the humic substance (2,000 mg L–1) was made in 50 mM NaOH.

Procedure A flow chart for the complete sample treatment is shown in Fig. 1. As a sample for a recovery experiment, 1 L of 10 mM KH2PO4 was mixed with 5 mL of a humic substance stock solution to give a concentration of 10 mg L–1, and this was spiked with 0.5 mL of the surrogates solution. For a quantitative experiment, a larger-volume (18 L) surface water sample was provided to obtain lower quantitation limits. The sample was spiked with 0.1 mL of SS. In addition, a commercial HA-fortified sample was also prepared by adding 27 mL of the HA stock solution (spiked concentration 3 mg L–1) and 0.1 mL of SS to sample 4. After shaking vigorously, the mixture was adjusted to pH 2 or 9 using 2 M HCl or 2 M NaOH and left overnight at room temperature to achieve sufficient aggregation of HA. The equilibrated mixture was first filtered through a glass fiber filter, and then percolated with a preconditioned membrane. After the entire sample was drawn through the filter, the par-

1856 Table 1 GC-MS analytical conditions Column

BPX-DXN

RH-12 ms

GC oven temperature program Injection temperature (°C) Initial temperature (°C) Initial time (min) Rate 1 (°C min–1) Final temperature 1 (°C) Rate 2 (°C min–1) Final temperature 2 (°C) Final time (min)

290 100 1.5 20 210 3 315 5

290 100 1.5 20 210 3 290 5

MS Resolution Ion current (µA) Electron voltage (eV) Ion source temperature (°C)

ca. 10,000 550 38 290

280

gate was calculated by comparing its peak area on the SIM chromatogram with that of the recovery standard.

Results and discussion Effects of humic substances on recovery of analytes from water by SPE

Fig. 1 Flow chart for the complete process for determination of PCDD/Fs and CoPCBs in water. aA humic substance was dissolved in 10 mM KH2PO4; bsurrogates solution; cextraction standard solution; d50 µL for a recovery test and 20 µL for a quantitative test; erecovery standards solution ticulate matters (PM) deposited on the bottom of the sample bottles were rinsed with water (for pH 9 samples) or 10 mM HCl (for pH 2 samples) and transferred onto the filter. Thereafter, for the recovery test, the glass filter and the membrane were separately extracted by PLE to evaluate the recovery from each part. With regard to the quantitative test, the filters and the membranes were put together, fortified with 10 µL of ESS, and extracted by PLE. Each extract was cleaned up by means of multilayer column and alumina column chromatography according to the procedure in Fig. 1. Quantification by HRGC-HRMS An aliquot (1 µL) of a resulting solution was injected into the GC, equipped with a BPX-DXN column for analysis from PCDD/Fs (60 m×0.25-mm i.d.) and a RH-12 ms column for analysis of CoPCBs (60 m×0.25-mm i.d.), in the splitless mode. Acquisition parameters for selected ion monitoring (SIM) mode followed those given elsewhere [13]. Other GC/MS conditions are shown in Table 1. The mass profiles of the selected ions were obtained during GC elution. Identification was based on examination of isotopic ratios of M+ and (M+2)+ or (M+2)+ and (M+4)+, and the GC retention time. The area of the mass profile peaks of the quantitative ions was used for the quantitative analysis. The quantitation was performed using an isotope dilution method. Recovery of the surro-

Association of PCDD/Fs and CoPCBs with humic substances increases their concentrations in filtered water [14]. Therefore, optimization of the extraction procedure from the dissolved layer is required for exact measurement of the target analytes in humic acid containing water. As mentioned in the “Introduction”, dioxins, similarly to other hydrophobic substances, are believed to associate strongly with humic materials. Therefore, effective capture of the analyte–humic associate is essential for sufficient recovery of the surrogates. Pichon et al. have indicated in their report on pesticide residues analysis that acidification of water samples promotes co-extraction of HA with a C18 cartridge [15]. On the basis of their findings, we examined the effect of acidification on recovery of spiked analytes from humic material solutions. Recovery charts obtained for humic acid solution spiked with 13C-labeled compounds are illustrated in Figs. 2 and 3. In the recovery experiments using the HA standard and the commercial HA, the overall recovery of 13C-labeled compounds on processing at pH 9 was insufficient (<60%), and especially poor for higher-chlorinated PCDD/Fs (15– 30%). On the other hand, processing at pH 2 gave excellent results (>70% of recovery) for all compounds. Since all analytes are nonionic compounds, the recovery should be dependent on the behavior of HA–analyte associates. When a surrogates-spiked commercial HA solution was processed at pH 2, recovery rates of the 13C-labeled compounds from the filter and the membrane were 33–46% and 45–54%, respectively (Fig. 2B). The compounds recovered from the filter are likely to be present in the HA aggregates. On the other hand, based on the results of our

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Fig. 2A, B Recovery of 13C-labeled compounds from spiked commercial humic acid solution by prefiltration and SPE at pH 9 (A) or 2 (B). The interval on top of each bar chart indicates mean±standard deviation of the summed recovery rate for triplicate determinations

additional experiment, it is assumed that those recovered from the membrane were also present as associates with HA molecules. Rinsing the membrane with water after percolation markedly lowered the recovery of the compounds from the membrane (data not shown). It is likely that target-associated HA molecules which are not aggregated at pH 2 are retained on the C18 membrane in their nonionic form. When the membrane is rinsed with neutral water, the retained associates probably change to their ionic form, leading to their desorption from the membrane. Quantitative analysis of acidified surface water by SPE The results obtained from the recovery experiments suggest that HA in surface water may be responsible for the insufficient recovery of the surrogates. As mentioned above, HA–surrogate associates are regarded as behaving

Fig. 3A, B Recovery of 13C-labeled compounds from spiked humic acid standard solution by prefiltration and SPE at pH 9 (A) or 2 (B). The interval on top of each bar chart indicates mean±standard deviation of the summed recovery rate for triplicate determinations

as follows when a water sample is acidified and processed: (i) they are aggregated and are retained on the filter, (ii) they change to their nonionic form and are retained on the membrane. Based on these assumptions, a conventional analytical method [3] was modified for improvement of recovery of the surrogates. First, a water sample was adjusted to pH 2 and equilibrated overnight to allow sufficient aggregation of HA–analyte associates. Second, all devices and materials (sample bottles, filtration device, filters, and C18 membranes) were rinsed with 10 mM HCl instead of water after processing to prevent redissolution or desorption of the associates. Four surface water samples were subjected to determination of PCDD/Fs and CoPCBs using the modified method, and the recovery of the surrogates and quantitative values was compared with those at pH 9 (since the pH values of all the samples used in this study were in the range of 8.5–9.0, unadjusted samples were regarded as pH 9 samples). In addition, Sample 4 fortified with the commercial HA (3 mg L–1 sample) was also determined.

1858 Table 2 Quantitation of PCDD/Fs and CoPCBs in surface water samples by SPE pH DOC (mgC L–1)a

Sample 1 8.8 11.1 pH 9

Recovery of labeled compounds (%) TeCDD1 72 PeCDD1 69 HxCDD1 57 HxCDD2 55 HxCDD3 56 HpCDD1 50 OCDD 55 TeCDF1 74 PeCDF1 69 PeCDF2 71 HxCDF1 56 HxCDF2 56 HxCDF3 55 HxCDF4 58 HpCDF1 52 HpCDF2 52 OCDF 50 1,2,3,4,6,7-HxCDD 94 (extraction spike) TeCB1 80 TeCB2 75 PeCB1 66 PeCB2 68 PeCB3 72 PeCB4 71 PeCB5 65 HxCB1 65 HxCB2 64 HxCB3 67 HxCB4 64 HpCB1 65 Concentration (pg L-1) TeCDD1 PeCDD1 HxCDD1 HxCDD2 HxCDD3 HpCDD1 OCDD TeCDF1 PeCDF1 PeCDF2 HxCDF1 HxCDF2 HxCDF3 HxCDF4 HpCDF1 HpCDF2 OCDF TeCB1 TeCB2 PeCB1 PeCB2

<0.01 0.046 0.050 0.18 0.12 1.7 9.8 0.054 0.075 0.14 0.12 0.12 0.16 0.24 0.46 0.071 0.44 3.0 0.11 11 1.3

Sample 2 8.5 11.1

Sample 3 9.0 10.9

Sample 4 8.7 13.2

pH 2

pH 9

pH 2

pH 9

pH 2

pH 9

+HAb pH 9

+HAb pH 2

89 94 84 89 93 84 79 86 92 93 88 87 90 84 86 80 81 106

87 81 73 74 67 61 59 89 82 84 68 69 69 68 59 64 60 101

89 92 94 93 87 85 95 93 100 94 90 92 87 91 89 90 95 105

92 80 80 77 75 63 55 82 75 79 82 74 81 68 65 62 58 117

96 86 89 90 90 86 78 95 89 88 96 89 97 88 85 89 83 108

60 59 52 51 48 44 45 64 56 58 49 49 47 53 45 45 43 90

27 23 21 22 23 21 26 27 24 24 22 22 22 23 23 20 24 90

86 78 87 86 77 75 83 89 83 82 86 87 86 87 78 78 83 105

93 90 88 87 86 88 88 94 94 88 97 93

87 80 86 90 86 86 84 79 81 79 85 80

92 90 88 96 92 92 104 92 95 93 98 102

82 87 70 80 80 79 77 76 72 70 75 68

97 94 93 99 97 96 100 99 100 96 111 108

74 74 67 67 72 68 62 57 57 59 55 50

35 37 34 37 29 38 31 31 36 34 33 33

80 78 81 82 70 82 79 76 80 78 77 75

0.032 0.11 0.14 0.16 0.16 1.7 13 0.080 0.094 0.20 0.16 0.16 0.062 0.29 0.68 0.11 0.84 4.0 0.25 15 1.0

<0.01 0.073 0.090 0.18 0.18 2.1 12 0.077 0.10 0.21 0.17 0.16 0.064 0.25 0.67 0.091 0.75 0.76 0.10 1.8 0.22

<0.01 0.13 0.20 0.44 0.36 4.0 20 0.15 0.19 0.46 0.31 0.30 0.12 0.50 1.2 0.19 1.8 1.0 0.13 2.3 0.31

<0.01 0.034 0.055 0.14 0.090 1.1 6.8 0.038 0.059 0.14 0.13 0.090 0.060 0.28 0.30 0.065 0.31 2.9 0.17 10 1.2

0.017 0.058 0.071 0.11 0.094 1.2 13 0.069 0.067 0.15 0.13 0.15 0.051 0.26 0.69 0.081 0.84 5.4 0.29 18 1.4

0.012 0.043 0.065 0.080 0.086 1.0 8.1 0.055 0.063 0.13 0.12 0.12 0.055 0.22 0.51 0.078 0.54 5.1 0.28 17 1.4

0.045 0.076 0.14 0.17 0.14 2.0 19 0.078 0.087 0.19 0.17 0.18 0.080 0.33 0.80 0.13 1.2 4.5 0.25 15 1.0

<0.01 0.068 0.082 0.19 0.17 1.7 7.9 0.073 0.10 0.17 0.14 0.14 0.065 0.23 0.53 0.080 0.51 0.92 0.11 1.9 0.28

1859 Table 2 (continued) pH DOC (mgC L–1)a

PeCB3 PeCB4 PeCB5 HxCB1 HxCB2 HxCB3 HxCB4 HpCB1 Sum TeCDDs Sum PeCDDs Sum HxCDDs Sum HpCDDs Sum TeCDFs Sum PeCDFs Sum HxCDFs Sum HpCDFs Total WHO-TEQ (pgTEQ L–1) aDissolved

Sample 1 8.8 11.1

Sample 2 8.5 11.1

Sample 3 9.0 10.9

Sample 4 8.7 13.2

pH 9

pH 2

pH 9

pH 2

pH 9

pH 2

pH 9

+HAb pH 9

+HAb pH 2

22 0.41 0.20 1.6 0.44 0.36 0.037 0.10 3.5 4.0 2.4 3.3 1.8 1.6 1.5 0.8 0.27

23 0.59 0.16 1.3 0.42 0.34 0.039 0.11 2.5 3.7 1.9 2.2 1.7 1.6 1.3 0.6 0.23

35 1.0 0.24 2.1 0.75 0.87 0.044 0.17 3.0 1.0 1.3 2.6 2.4 1.6 1.7 1.2 0.30

34 0.85 0.22 2.1 0.69 0.80 0.049 0.13 3.1 1.1 1.2 2.0 2.5 1.8 1.4 1.0 0.25

29 0.63 0.32 2.5 0.75 1.0 0.059 0.26 5.2 1.6 2.0 4.1 2.7 2.3 2.1 1.6 0.42

27 0.69 0.28 2.4 0.69 0.90 0.064 0.17 6.0 1.8 2.0 3.5 2.7 2.4 1.9 1.3 0.43

3.4 0.11 0.24 0.49 0.22 0.24 0.069 0.19 2.6 2.0 3.4 4.6 2.4 2.0 1.7 1.1 0.35

4.8 0.27 0.34 0.76 0.28 0.27 0.13 0.20 4.8 4.2 7.2 8.5 4.3 3.9 3.2 2.1 0.71

3.6 0.15 0.26 0.56 0.17 0.17 0.07 0.17 2.4 2.2 3.4 3.7 2.5 1.7 1.5 0.9 0.33

organic carbon (concentration of total organic carbon in the filtrate) humic acid was added (3 mg L–1 sample) before adjustment of sample pH

bCommercial

Table 3 Comparison of quantitation factor and extraction factor for OCDD Sample 1

Quantitation factora Extraction factorb

Sample 2

Sample 3

Sample 4

pH 9

pH 2

pH 9

pH 2

pH 9

pH 2

pH 9

+HAc pH 9 +HAc pH 2

0.81 0.72

0.54 0.68

1.3 1.3

0.78 1.2

1.9 1.8

1.3 1.8

0.99 0.62

1.6 0.66

0.63 0.66

aPeak

area ratio of native OCDD to 13C-OCDD area ratio of native OCDD to extraction standard (13C-1,2,3,4,6,7-HxCDD) cCommercial humic acid was added (3 mg L–1 sample) before adjustment of sample pH bPeak

The results obtained are shown in Table 2. Insufficient recovery of higher-chlorinated CDD/Fs (HpCDD/Fs and OCDD/F) surrogates (43–65%) was obtained for all samples when the sample was processed at pH 9, but this insufficient recovery was improved by acidification of the sample. In regard to quantitative values, these results did not agree with the recovery results: the quantitative values of higher-chlorinated CDD/Fs at pH 9 were apparently higher than those obtained at pH 2 (Table 2). In Table 3, the extraction factor (native OCDD peak area ratio to 13C-1,2,3,4,6,7-HxCDD) and the quantitation factor (native OCDD area ratio to 13C-OCDD) at pH 2 are compared with those at pH 9. For each sample, the quantitation factor shows an apparent difference between at pH 2 and at pH 9, whereas the extraction factor did not indicate a significant difference. Therefore, for higher-chlorinated PCDD/Fs, the extraction factors of native compounds would not be as sensitive to pH change as recovery of the surrogates. This would induce a systematic overestimation of their concentration at pH 9 and is the reason for the relative poorer recovery of surrogates at this pH.

These results suggest a difference between the behavior of the target analytes and that of the surrogates in the water samples. We conclude, on the basis of our recovery results, that during processing at pH 9, small quantities of the spiked surrogates associate with dissolved HA (as a solution-phase phenomenon), being carried through the C18 membrane. When the sample was processed at pH 2, however, the associates appear to be retained on the filter or the membrane. On the other hand, it is likely that, regardless of the pH of the sample, most of the native higher-chlorinated CDD/Fs were adsorbed onto PM and trapped in the filter. The quantitation of the HA-fortified sample 4 exhibited the remarkable tendency of HA to lower the recovery of the surrogates and to cause resultant overestimation of the quantitative values. The results in Table 2 do not provide evidence of a correlation between DOC concentration and the recovery rates of the surrogates. It is well known that FA is a major component of DOC in ordinary surface water [5]. We also examined a recovery experiment using the FA standard and confirmed that it does not influence the recovery of

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the surrogates (data not shown). On the other hand, the addition of HA to a water sample resulted in poor recovery of the surrogates and overestimation of the quantitative values for most of the homologues (Tables 2 and 3). Thus, it appears that the behavior of the surrogates in surface water samples is dependent on the concentration of HA which can associate with them in the dissolved organic matters, but not on the total DOC concentration. In this study, a commercial HA and a HA standard derived from soil were used because we were unable to obtain aquatic HA. Since the former’s molecular characteristics are different from those of aquatic HA [16], the obtained recovery results cannot be applied to water samples. However, aquatic HA is likely to have the same basic properties (i.e., interaction with nonpolar dioxins and nonionization under acidic conditions) as commercial or soil HA. The results in Table 2 (improvement of recovery by acidification of the sample) are consistent with those for HA, and therefore suggest that HA in surface water is likely to be responsible for insufficient recovery of the surrogates. When carrying out determination by isotope dilution, a major premise is that the native target analyte and its surrogate behave in a closely similar manner. Our results, however, indicate that an insufficiently recovered surrogate cannot behave in the same manner as the corresponding native target analyte in analysis of PM and HA-containing water. In such cases, acidification of the water sample prior to SPE will give better recovery of the surrogates and more accurate quantitative values for the target analytes. Acknowledgements We thank Dr M Shinomiya (National Environmental Research and Training Institute, Ministry of the Envi-

ronment) for help in measuring the DOC concentration in the water samples.

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