摘要:

Speciation of Arsenic Animal Feed Additives by Microbore High-performance Liquid Chromatography with Inductively Coupled Plasma Mass Spectrometry Spiros A. Pergantis*†, Edward M. Heithmar and Thomas A. Hinners US Environmental Protection Agency, National Exposure Research Laboratory, Environmental Sciences Division, P.O. Box 93478, Las Vegas, NV 89193-3478, USA Phenylarsonic compounds have been used as poultry and swine feed additives for the purpose of growth promotion and disease prevention.Owing to the lack of suitable analytical methods, however, knowledge of their metabolism, environmental fate and impact remains incomplete. In order to compensate for this, analytical procedures were developed that allow the speciation of arsenic animal feed additives by using microbore high-performance liquid chromatography (mHPLC) coupled on-line with ICP-MS. More specifically, reversed-phase (RP) chromatographic methods were optimised to achieve the separation of various phenylarsonic acids from each other and from the more toxic inorganic arsenic compounds.This mode of chromatography, however, exhibits limitations, especially in the presence of naturally occurring organoarsenic compounds. The application of RP ion-pairing chromatography eliminates such shortcomings by minimising the co-elution of arsenic species. In general, the mHPLC–ICP-MS methods developed in this study provide high selectivity, extremely good sensitivity, low limits of detection (low-ppb or sub-pg amounts of As), require small sample volumes ( < 1 ml), minimise waste and operate most efficiently under low mobile-phase flow rates (15–40 ml min21), which are compatible for use with other types of mass spectrometers, e.g., electrospray.Reference materials containing naturally occurring arsenic compounds were spiked with phenylarsonic compounds and then analysed by using the procedures developed in this study. Keywords: Arsenic; speciation; inductively coupled plasma mass spectrometry; microbore high-performance liquid chromatography; reference materials; animal feed additives A number of phenylarsonic compounds have been shown to control cecal coccidiosis in poultry, and also act as growth promoters, providing improved feed conversion, better feathering and increased egg production and pigmentation. 4-Hydroxy- 3-nitrophenylarsonic acid (roxarsone), p-arsanilic acid (p-ASA), 4-nitrophenylarsonic acid (4-NPAA), p-ureidophenylarsonic acid (p-UPAA) and benzenearsonic acid have all been used for such purposes and, with the exception of p- UPAA, are still in use today. Variation in the substituents on the aromatic ring results in differences in the growth-promoting and disease-controlling effects of the compounds.Thus, roxarsone and p-ASA are approved as animal feed additives for both poultry and swine, whereas 4-NPAA and p-UPAA are approved only for controlling blackhead disease in turkeys.1–3 Even though studies pertaining to the metabolism,4 toxicity5,6 and excretion of these compounds have been conducted, their physiological role, environmental fate and impact are still not well understood.In order to improve our understanding of these processes, it is required that analytical methods be developed that will allow for the speciation of phenylarsonic compounds and their metabolites. So far, only a limited number of methods have been developed and used for the determination of this class of compounds. Most of these methods are for the targeted analysis of arsenic animal feed additive compounds, and are therefore not necessarily suitable for the detection of potential metabolites.For example, gas chromatography with flame ionization detection, which has been used for the determination of p-UPAA and p-ASA, is not suitable for the determination of roxarsone and is also interference prone.7 A spectrophotometric method for the determination of p-UPAA in the animal feed additive Carbasone has also been reported.8 This method involves a coupling reaction with N-1-naphthylethylenediamine.The colored product that forms is extracted with butanol and subsequently measured photometrically; again, interferences can cause difficulties with this determination. Thin-layer chromatography has been used for the separation and identification of roxarsone, p-ASA, 4-NPAA and p-UPAA; chromogenic reagents were used for detection.9 HPLC has also been used to separate some arylarsenicals, but not specifically those used as animal-feed additives.10–12 Also, an LC method has been developed for the detection and quantification of roxarsone in poultry feed.The drug is extracted by means of a phosphate buffer and determined using solid-phase extraction in combination with reversed-phase (RP) LC with UV detection.13 HPLC coupled on-line with ICP-MS has been used for the determination of roxarsone14,15 and other arsenic animal feed additive compounds.14 More recently, a number of MS methods have been tested as a means to characterize structurally arsenic animal feed additives.16–18 In this paper we report on the development of microbore HPLC (mHPLC)–ICP-MS methods, which allow the speciation of arsenic animal feed additives present in environmental and biological samples.Of primary interest was the development of methods that allow us to differentiate between arsenic animal feed additives and naturally occurring arsenic compounds, and also provide information concerning the presence of potential metabolites.Experimental Instrumentation A VG PlasmaQuad II STE ICP-MS system (VG Elemental, Winsford, Cheshire, UK) was used for elemental detection. The quadrupole mass analyser was operated in the single-ion monitoring mode (m/z 75) for the determination of arsenic. The microscale flow injection (mFI) and mHPLC set-ups have been described in detail in previous work.14 A 0.5 or 1 ml internal loop injector (Valco Instruments, Houston, TX, USA) was used for both mFI and mHPLC.A Model 100DM syringe pump (Isco, Lincoln, NE, USA) was used to deliver microflows between 15 and 80 ml min21. † Present address: Department of Chemistry, Birkbeck College, University of London, Gordon House, 29 Gordon Square, London, UK WC1H 0PP. Analyst, October 1997, Vol. 122 (1063–1068) 1063Chemicals and Reagents Inorganic arsenic standard solutions used for the mFI–ICP-MS experiments were prepared by diluting 1000 ppm standards (Inorganic Ventures, Lakewood, NJ, USA) with de-ionized water (18 MW) acidified to 0.05% v/v with doubly distilled nitric acid (Seastar, Sidney, BC, Canada).The mFI carrier was also 0.05% nitric acid. Although 1% nitric acid is more commonly used in ICP-MS, a lower acid concentration was used in this study to avoid possible corrosion of the stainlesssteel syringe pump. No problems with memory effects were caused by the use of 0.05% nitric acid carrier.To minimise contributions to the blanks from leached pump materials, the small-diameter steel tubing and valves on the pump were replaced with Teflon hardware. The following certified reference materials were analysed for their arsenic species by using mHPLC–ICP-MS: Trace Elements in Water [National Institute of Standards and Technology (NIST) Standard Reference Material (SRM) 1643a]; Trace Metals in Drinking Water (High-Purity Standards, Charleston, SC, USA); Toxic Metals in Urine (NIST SRM 2670n); and San Joaquin Soil (NIST SRM 2709).Soil extractions were performed by using 0.3 m H3PO4 and sonicating for 60 min. All solutions were filtered with 0.45 mm syringe filters (nylon Acrodisc, Gelman Sciences, Ann Arbor, MI, USA) and analysed without further dilution. The separation of arsenic compounds by using RP-mHPLC was accomplished using a mobile phase consisting of 0.1% trifluoroacetic acid (Sigma, St.Louis, MO, USA) and 5–10% v/v methanol in water. For the RP ion-pairing (IP) mHPLC separations, tetrabutylammonium hydroxide (Alfa Products, Danvers, MA, USA) was used as the ion-pairing reagent at concentrations ranging from 1 to 5 mm. Malonic acid (Sigma) was used to adjust the mobile phase pH. The analytical mHPLC column, used for both RP and RP-IP chromatography, was a 150 mm 3 1 mm id stainless-steel column packed with Spherisorb 3 mm C18 material (Isco).HPLC mobile phases were filtered and de-gassed using a 35 mm all-glass filter holder (Millipore, Bedford, MA, USA), fitted with a 0.45 mm hydrophilic nylon filter (Cuno, Meriden, CT, USA) for aqueous solvents and a 0.5 mm PTFE filter (Cole-Parmer, Chicago, IL, USA) for organic solvents. Chromatographic standards were prepared from 1000 mg l21 As aqueous solutions of the following arsenic compounds: parsanilic acid (p-ASA) (Eastman Organic Chemicals, Rochester, NY, USA), 3-nitro-4-hydroxyphenylarsonic acid (roxarsone) (ICN Biochemicals, Cleveland, OH, USA), 4-nitrophenylarsonic acid (4-NPAA) (Aldrich, Milwaukee, WI, USA), dimethylarsinic acid (DMA) (Sigma), 4-hydroxyphenylarsonic acid (4-OH) (Eastman Kodak), disodium methylarsonate (MMA) (Chemical Service, West Chester, PA, USA), sodium meta-arsenite (Aite) (Sigma), and sodium arsenate (Aate) (Sigma).Procedures Time resolved acquisition (TRA) software provided by VG was used to acquire data during mHPLC–ICP-MS experiments.Data files of response versus time were exported as ASCII files and analysed using commercial spreadsheet software. The mFI– and mHPLC–ICP-MS response was optimised for the detection of arsenic at m/z 75 using carrier flow rates of 80 and 40 ml min21, respectively. The optimisation was carried out by employing a 60 ml loop fitted on a ceramic injector (SVI- 6U7) (Analyticon Instruments, Springfield, NJ, USA). The large volume loop provided quasi-continuous sample introduction, which lasted long enough for signal optimisation to be achieved.For the HPLC optimisation a 60 ml loop was connected post-column; this provided a signal equivalent to that obtained in the continuous flow mode. All instrument parameters were varied iteratively to reach the apparent optimum response. Results and Discussion Separation of Arsenic Animal Feed Additives Using RP- mHPLC–ICP-MS RP-mHPLC methods were developed and used for the separation of four phenylarsonic acids and two inorganic arsenic compounds.A mobile phase consisting of 0.1% trifluoroacetic acid (TFA) in water, with a variable percentage (5–15% v/v) of methanol, was used for their separation. The mHPLC stationary phase consisted of silica based C18 material (3 mm particle diameter, 12% carbon loading). The chromatograms obtained are presented in Fig. 1. Present in the injected sample was 4-hydroxyphenylarsonic acid (4-OH). Even though this compound has not been used as an animal feed additive, it may prove to be a decomposition product or metabolite of roxarsone or may adequately serve as a chromatographic internal standard.As mentioned previously, the RP-mHPLC method developed allows for the separation of inorganic arsenic species from various of the phenylarsonic acids. This separation is of particular importance, primarily because of the relatively high toxicity of the inorganic arsenic species. Thus, the RP chromatographic method can be used to monitor arsenic animal feed additives for the presence of inorganic arsenic impurities, and also to detect possible degradation of the phenylarsonic acids to their respective inorganic forms.As shown in Fig. 1, the extensive retention of the NO2-containing arsenicals (roxarsone and 4-NPAA) by the LC stationary phase results in excessive Fig. 1 RP-mHPLC–ICP-MS traces for six arsenic compounds. The mobile phase consisted of water (0.1% TFA) and 5–15% methanol, applied at a flow rate of 40 ml min21. 1064 Analyst, October 1997, Vol. 122peak tailing. A larger amount of methanol in the mobile phase reduces peak tailing significantly, but also provides poorer overall resolution. In addition, the methanol content affects the sensitivity of the ICP-MS detector. Signal enhancement for arsenic in the presence of organic solvents has been observed and reported extensively by others.19,20 This occurrence has major implications, especially when using gradient-elution methods which make use of variable amounts of organic solvents in conjunction with HPLC–ICP-MS.It has been proposed that the presence of organic solvents in the plasma alters the energetics of the ICP. Even when the introduction of organic solvent into the plasma is discontinued, a relatively long period of time is required before the ICP-MS sensitivity returns to its pre-solvent level. To explore further the possible use of RP chromatography for the speciation of phenylarsonic acids in environmental samples, we investigated the effects of naturally occurring arsenicals on the RP chromatograms.The RP-mHPLC separation of various phenylarsonic acids (arsenic animal feed additives) from naturally occurring arsenic compounds is presented in Fig. 2. These chromatograms were obtained with no methanol present in the mobile phase. Less efficient separations occurred in the presence of even small amounts of methanol ( < 0.5%). The mobile phase flow rate also had a pronounced effect on separation efficiency. The best separation was obtained at the lowest-tested flow rate (15 ml min21). The main drawback, however, was the incomplete separation of DMA from p-ASA. In addition, excessive peak broadening occurred in the low flow rate regime ( < 30 ml min21).Flow rates > 55 ml min21 were not considered, mainly because of the resulting high back-pressure ( > 3000 psi), which could potentially damage the mHPLC column.It should also be mentioned that when the mobile phase does not contain any methanol, the phenylarsonic acids roxarsone and 4-NPAA do not elute from the HPLC column. Thus, a preliminary chromatographic run using a mobile phase containing 5–15% methanol must be conducted in order to investigate the presence of the two nitro-containing arsenicals (roxarsone and 4-NPAA). Separation of Arsenic Animal Feed Additives Using RP-IP- mHPLC–ICP-MS Because RP chromatography provides poor separation of the arsenic animal feed additives from some of the naturally occurring arsenic compounds (Aite, Aate, MMA and DMA), alternative chromatographic modes of separation were investigated and further optimised to provide efficient separations.To accomplish this, RP-IP-mHPLC was further investigated. To assist in reaching the optimum conditions required for such separations, a plot of the apparent charge (Qapp) of each of the arsenic compounds as a function of mobile phase pH was constructed (Fig. 3). This plot provides a qualitative estimate of the extent of interaction of the anion-pairing reagent with the arsenic-containing analyte under various pH conditions. Qapp for arsenate was calculated as follows: Qapp AsO HAsO H AsO AsO HAsO H AsO H AsO = - + + + + + - - - - - - 3 2 4 3 4 2 2 4 4 3 4 2 2 4 3 4 [ ] [ ] [ ] [ ] [ ] [ ] [ ] The Qapp values for the other arsenic compounds were derived similarly.The pKa values21 used for these calculations were as follows: pKa[Aite] = 9.2, pKa[Aate] = 2.2, 6.98, 11.5, pKa[DMA] = 1.3,22 6.3, pKa[MMA] = 3.41, 8.18, pKa[p- ASA] = 2, 4.02, 8.92. The chromatogram presented in Fig. 4, obtained under RPIP- mHPLC conditions, exhibits sufficient separation of p-ASA Fig. 2 RP-mHPLC–ICP-MS traces for p-ASA (D) and 4-OH (E) in the presence of naturally occurring arsenic compounds: arsenite and arsenate (A), methylarsonic acid (B) and dimethylarsinic acid (C).The mobile phase consisted of water (0.1% TFA). The flow rates varied between 15 and 55 ml min21. Fig. 3 Apparent charge of arsenic compounds as a function of solution pH. Arrows indicate normal operating range of pH for reversed-phase anionpairing chromatography of arsenic acids. Fig. 4 RP-IP-mHPLC–ICP-MS traces for five arsenic compounds. The mobile phase consisted of 1 mm tetrabutylammonium hydroxide and 0.5% methanol in de-ionised water (pH 5.28), applied at a flow rate of 40 ml min21.Analyst, October 1997, Vol. 122 1065from four other arsenic compounds which have been found to occur naturally in various environmental samples. The elution order of these compounds at pH 5.3 is in agreement with that predicted by the Qapp versus pH plot (Fig. 3). A compound with high Qapp is expected to interact to a greater extent with the tetrabutylammonium ion-pairing reagent. Stronger interaction results in longer retention times. However, it should be noted that factors other than pH significantly influence the chromatographic behavior of the arsenic compounds.The most important of these factors include the ion-pairing reagent concentration and the methanol content of the mobile phase. Separation of p-ASA from Arsenic Compounds Present in Reference Materials Initially the effects of large amounts of NaCl on the chromatographic separation of arsenic compounds were investigated. The resulting chromatograms from these experiments are presented in Fig. 5. It was observed that the presence of 0.1% NaCl in the sample had a significant effect on the chromatographic separations (retention times, peak shapes), particularly when a low ion-pairing concentration (1–2 mm TBAH) was used. In order to minimise this problem, which is especially pronounced when speciating arsenic in biological materials such as urine, 5 mm TBAH was used. Also, the mobile phase pH was adjusted to allow for the separation of chloride from the other arsenic compounds, thus eliminating potential interferences caused by ArCl+.Freeze-dried urine (NIST SRM 2670) containing ‘normal levels’ of arsenic (as indicated by NIST) was analysed for its arsenic content. The freeze-dried urine sample was reconstituted with de-ionized water and then spiked with p-ASA and 4-OH. The resulting chromatogram is shown in Fig. 6. Resolution between DMA and MMA was sacrificed by appropriately adjusting the mobile phase pH in order to achieve improved separation between the Cl2 and arsenate species present.The determination of arsenate is of considerable importance mainly because its toxicity is substantially higher than that of MMA or DMA. Good separation of 4-OH, used as an internal standard in this case, was also accomplished. It should be noted that the original (non-spiked) urine SRM did not contain any measurable amounts of p-ASA or 4-OH. It should also be noted that the RP anion-pairing chromatography used in this study is not capable of differentiating between arsenobetaine and arsenite, as these two compounds co-elute.Also, there exists the possibility that other non-retained arsenic species may co-elute in this front-end peak. Arsenobetaine is normally present in human urine following the consumption of seafood.23 Previous investigations using this particular reference material have shown that indeed the first peak is the result of arsenite and arsenobetaine.18,24 Solutions referred to as Trace Elements in Water (NIST SRM 1643a) and Trace Metals in Drinking Water (High-Purity Standards) were also spiked with p-ASA and 4-OH, and subsequently analysed for their content of arsenic species (Fig. 7).The spiked arsenicals were separated from arsenite and arsenate, which were originally present in the water reference materials. Furthermore, San Joaquin Soil (NIST SRM 2709) was extracted using 0.3 m H3PO4 and then spiked with p-ASA and 4-OH.The resulting chromatogram is presented in Fig. 8. The soil extract, prior to spiking, was found to contain only arsenate as the single arsenic compound present. The limits of detection for each arsenic compound, achieved under various modes of chromatography, are summarised in Table 1. Because of the extremely high sensitivity of the ICPMS detector, the chromatographic methods described are eminently suitable for the development of methodologies for the determination of arsenic animal feed additives in real samples.These methods are particularly suited for the analysis of samples containing limited amounts of analyte. mFI–ICP-MS of Arsenic Animal Feed Additives In previous work, a mFI–ICP-MS technique was developed and applied to the determination of total arsenic.14 Of interest in this study was to investigate further any sensitivity variations observed when analysing arsenic animal feed additives by using mFI–ICP-MS. A number of reports have claimed sensitivity variations for other organoarsenic compounds determined using continuous-flow ICP-MS, particularly for arsenicals present in marine organisms.20 So far, no sensitivity data have been reported regarding arsenic animal feed additives.Fig. 9 shows the peaks obtained for the four phenylarsenical compounds discussed in this paper. Each peak corresponds to an Fig. 5 Effect of salt and ion-pair reagent concentrations on the separation of four arsenic compounds.Fig. 6 RP-IP-mHPLC–ICP-MS trace for urine (NIST SRM 2670n) spiked with p-ASA and 4-OH. The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. 1066 Analyst, October 1997, Vol. 122injection of 80 pg of As (1 ml injections of a phenylarsenical solution containing 80 ppb of As). The fact that no sensitivity variations were observed permits the use of mFI–ICP-MS for the quantification of arsenic animal feed additives regardless of which phenylarsonic species are present in the sample and without their prior conversion into a common species by employing some form of digestion.Conclusions We have demonstrated the development and successful application of a variety of chromatographic methods for the identification of arsenic animal feed additives. More specifically, RP- mHPLC was used for the separation of arsenic animal feed additives from inorganic arsenic species. This mode of chromatography, however, did not allow the complete separation of arsenic animal-feed additives from naturally occurring organoarsenic compounds.RP–IP-mHPLC was used successfully for this purpose. Furthermore, the use of mHPLC offers the advantage of operating at very low flow rates (15–40 ml min21), thus allowing for minimisation of waste and the use of small sample volumes, and also offers flow rate compatibility with other mass spectrometric detectors such as electrospray and continuous-flow fast atom bombardment mass spectrometry.The latter feature may prove to be a great benefit for the structural identification of metabolites or decomposition products for which synthetic standards are not yet available. The use of the ICP-MS detector offers additional benefits, such as high selectivity and extremely good sensitivity, for the speciation of trace levels of arsenic in samples of environmental and biological origin. The US Environmental Protection Agency (EPA), through its Office of Research and Development (ORD), funded the research described here.This paper has been subjected to the Agency’s peer review and has been approved as an EPA publication. The US Government has a non-exclusive, royaltyfree license in and to any copyright covering this paper. Mention of trade names or commercial products does not constitute Fig. 7 RP-IP-mHPLC–ICP-MS traces for A, water (NIST SRM 1643a) spiked with p-ASA and 4-OH and B, water (High-Purity Standards, Drinking Water) spiked with p-ASA and 4-OH.The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. Fig. 8 RP-IP-mHPLC–ICP-MS trace for soil extract (NIST SRM 2709 San Joaquin Soil) spiked with p-ASA and 4-OH. The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. Table 1 Limits of detection (defined as three times the standard deviation of the background) for arsenic compounds obtained under RP- and RP-IP-m- HPLC–ICP-MS conditions RP-IPRP- mHPLC– mHPLC– ICP-MS ICP-MS pg ml21 pg ml21 As pg As As pg As Arsenite (Aite) 0.10 0.10 0.6 0.6 Arsenate (Aate) 0.10 0.10 0.4 0.4 p-Arsanilic acid (p-ASA) 0.10 0.10 0.9 0.9 4-Hydroxyphenylarsonic acid (4-OH) 0.10 0.10 0.8 0.8 3-Nitro-4-hydroxyphenylarsonic acid (roxarsone) 0.12 0.12 n.d.* n.d. 4-Nitrophenylarsonic acid (4-NPAA) 0.26 0.26 n.d. n.d. * n.d.: not determined because the compounds do not elute from column under the specified conditions.Fig. 9 mFI–ICP-MS of arsenic animal feed additives. Each peak represents the injection of 1 ml of a solution of 80 pg ml21 As. Analyst, October 1997, Vol. 122 1067endorsement or recommendation for use. This work was performed while S.A.P. held a National Research Council/ CRD-LV Research Associateship. References 1 Gilbert, F. R., Wells, G. A. H., and Gunning, R. F., Vet. Rec., 1981, 109, 158. 2 Drugs Directorate, Health and Welfare Canada, Ottawa, Canada, 1981. 3 Compendium of Medicating Ingredients Brochures, Agriculture Canada, Ottawa, 5th edn., 1984. 4 Aschbacher, P. W., and Feil, V. J., J. Agric. Food Chem., 1991, 39, 146. 5 Edmonds, M. S., and Baker, D. H., J. Anim. Sci., 1986, 63, 553. 6 Rice, D. A., McMurray, C. H., McCracken, R. M., Bryson, D. G., and Maybin, R., Vet. Rec., 1980, 106, 312. 7 Weston, R. E., Wheals, B. B., and Kensett, M. J., Analyst, 1971, 96, 601. 8 Hoodless, R. A., and Tarrant, K. R., Analyst, 1973, 98, 502. 9 Morrison, J. L., J. Agric. Food Chem., 1968, 16, 704. 10 Maruo, M., Hirayama, N., Wada, H., and Kuwamoto, T., J. Chromatogr., 1989, 466, 379. 11 Hirayama, N., and Kuwamoto, T., J. Chromatogr., 1988, 457, 415. 12 Dodd, M., PhD Thesis, University of British Columbia, Vancouver, 1988. 13 Sapp, R. E., and Davidson, S., J. AOAC Int., 1993, 76, 956. 14 Pergantis, S. A., Heithmar, E. M., and Hinners, T. A., Anal. Chem., 1995, 67, 4530. 15 Dean, J. R., Ebdon, L., Foulkes, M. E., Crews, H. M., and Massey, R. C., J. Anal. At. Spectrom., 1994, 9, 615. 16 Pergantis, S. A., Cullen, W. R., Chow, D. T., and Eigendorf, G. K., J. Chromatogr. A, 1997, 764, 211. 17 Pergantis, S. A., Cullen, W. R., and Eigendorf, G. K., Biol. Mass Spectrom., 1994, 23, 749. 18 Pergantis, S. A., Winnik, W., and Betwoski, D., J. Anal. At. Spectrom., 1997, 12, 531. 19 Allain, P., Jaunault, L., Mauras, Y., Mermet, J.-M., and Delaporte, T., Anal.Chem., 1991, 63, 1497. 20 Larsen, E. H., and St�urup, S., J. Anal. At. Spectrom., 1994, 9, 1099. 21 Dean, J. A., in Lange’s Handbook of Chemistry, McGraw-Hill, New York, 13th edn., 1985. 22 Hansen, S. H., Larsen, E. H., Pritzl, G., and Cornett, C., J. Anal. At. Spectrom., 1992, 7, 629. 23 Le, X. C., Cullen, W. R., and Reimer, K. J., Clin. Chem., 1994, 40, 617. 24 Pergantis, S. A., Momplaisir, G.-M., Heithmar, E. M., and Hinners, T. A., in Proceedings of the 44th ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 12–16, 1996, American Society for Mass Spectrometry, East Lansing, MI, USA, p. 21. Paper 7/02691I Received April 21, 1997 Accepted July 14, 1997 1068 Analyst, October 1997, Vol. 122 Speciation of Arsenic Animal Feed Additives by Microbore High-performance Liquid Chromatography with Inductively Coupled Plasma Mass Spectrometry Spiros A. Pergantis*†, Edward M. Heithmar and Thomas A. Hinners US Environmental Protection Agency, National Exposure Research Laboratory, Environmental Sciences Division, P.O.Box 93478, Las Vegas, NV 89193-3478, USA Phenylarsonic compounds have been used as poultry and swine feed additives for the purpose of growth promotion and disease prevention. Owing to the lack of suitable analytical methods, however, knowledge of their metabolism, environmental fate and impact remains incomplete. In order to compensate for this, analytical procedures were developed that allow the speciation of arsenic animal feed additives by using microbore high-performance liquid chromatography (mHPLC) coupled on-line with ICP-MS.More specifically, reversed-phase (RP) chromatographic methods were optimised to achieve the separation of various phenylarsonic acids from each other and from the more toxic inorganic arsenic compounds. This mode of chromatography, however, exhibits limitations, especially in the presence of naturally occurring organoarsenic compounds.The application of RP ion-pairing chromatography eliminates such shortcomings by minimising the co-elution of arsenic species. In general, the mHPLC–ICP-MS methods developed in this study provide high selectivity, extremely good sensitivity, low limits of detection (low-ppb or sub-pg amounts of As), require small sample volumes ( < 1 ml), minimise waste and operate most efficiently under low mobile-phase flow rates (15–40 ml min21), which are compatible for use with other types of mass spectrometers, e.g., electrospray.Reference materials containing naturally occurring arsenic compounds were spiked with phenylarsonic compounds and then analysed by using the procedures developed in this study. Keywords: Arsenic; speciation; inductively coupled plasma mass spectrometry; microbore high-performance liquid chromatography; reference materials; animal feed additives A number of phenylarsonic compounds have been shown to control cecal coccidiosis in poultry, and also act as growth promoters, providing improved feed conversion, better feathering and increased egg production and pigmentation. 4-Hydroxy- 3-nitrophenylarsonic acid (roxarsone), p-arsanilic acid (p-ASA), 4-nitrophenylarsonic acid (4-NPAA), p-ureidophenylarsonic acid (p-UPAA) and benzenearsonic acid have all been used for such purposes and, with the exception of p- UPAA, are still in use today. Variation in the substituents on the aromatic ring results in differences in the growth-promoting and disease-controlling effects of the compounds. Thus, roxarsone and p-ASA are approved as animal feed additives for both poultry and swine, whereas 4-NPAA and p-UPAA are approved only for controlling blackhead disease in turkeys.1–3 Even though studies pertaining to the metabolism,4 toxicity5,6 and excretion of these compounds have been conducted, their physiological role, environmental fate and impact are still not well understood. In order to improve our understanding of these processes, it is required that analytical methods be developed that will allow for the speciation of phenylarsonic compounds and their metabolites.So far, only a limited number of methods have been developed and used for the determination of this class of compounds. Most of these methods are for the targeted analysis of arsenic animal feed additive compounds, and are therefore not necessarily suitable for the detection of potential metabolites.For example, gas chromatography with flame ionization detection, which has been used for the determination of p-UPAA and p-ASA, is not suitable for the determination of roxarsone and is also interference prone.7 A spectrophotometric method for the determination of p-UPAA in the animal feed additive Carbasone has also been reported.8 This method involves a coupling reaction with N-1-naphthylethylenediamine. The colored product that forms is extracted with butanol and subsequently measured photometrically; again, interferences can cause difficulties with this determination.Thin-layer chromatography has been used for the separation and identification of roxarsone, p-ASA, 4-NPAA and p-UPAA; chromogenic reagents were used for detection.9 HPLC has also been used to separate some arylarsenicals, but not specifically those used as animal-feed additives.10–12 Also, an LC method has been developed for the detection and quantification of roxarsone in poultry feed.The drug is extracted by means of a phosphate buffer and determined using solid-phase extraction in combination with reversed-phase (RP) LC with UV detection.13 HPLC coupled on-line with ICP-MS has been used for the determination of roxarsone14,15 and other arsenic animal feed additive compounds.14 More recently, a number of MS methods have been tested as a means to characterize structurally arsenic animal feed additives.16–18 In this paper we report on the development of microbore HPLC (mHPLC)–ICP-MS methods, which allow the speciation of arsenic animal feed additives present in environmental and biological samples.Of primary interest was the development of methods that allow us to differentiate between arsenic animal feed additives and naturally occurring arsenic compounds, and also provide information concerning the presence of potential metabolites. Experimental Instrumentation A VG PlasmaQuad II STE ICP-MS system (VG Elemental, Winsford, Cheshire, UK) was used for elemental detection.The quadrupole mass analyser was operated in the single-ion monitoring mode (m/z 75) for the determination of arsenic. The microscale flow injection (mFI) and mHPLC set-ups have been described in detail in previo work.14 A 0.5 or 1 ml internal loop injector (Valco Instruments, Houston, TX, USA) was used for both mFI and mHPLC. A Model 100DM syringe pump (Isco, Lincoln, NE, USA) was used to deliver microflows between 15 and 80 ml min21.† Present address: Department of Chemistry, Birkbeck College, University of London, Gordon House, 29 Gordon Square, London, UK WC1H 0PP. Analyst, October 1997, Vol. 122 (1063–1068) 1063Chemicals and Reagents Inorganic arsenic standard solutions used for the mFI–ICP-MS experiments were prepared by diluting 1000 ppm standards (Inorganic Ventures, Lakewood, NJ, USA) with de-ionized water (18 MW) acidified to 0.05% v/v with doubly distilled nitric acid (Seastar, Sidney, BC, Canada).The mFI carrier was also 0.05% nitric acid. Although 1% nitric acid is more commonly used in ICP-MS, a lower acid concentration was used in this study to avoid possible corrosion of the stainlesssteel syringe pump. No problems with memory effects were caused by the use of 0.05% nitric acid carrier. To minimise contributions to the blanks from leached pump materials, the small-diameter steel tubing and valves on the pump were replaced with Teflon hardware.The following certified reference materials were analysed for their arsenic species by using mHPLC–ICP-MS: Trace Elements in Water [National Institute of Standards and Technology (NIST) Standard Reference Material (SRM) 1643a]; Trace Metals in Drinking Water (High-Purity Standards, Charleston, SC, USA); Toxic Metals in Urine (NIST SRM 2670n); and San Joaquin Soil (NIST SRM 2709). Soil extractions were performed by using 0.3 m H3PO4 and sonicating for 60 min.All solutions were filtered with 0.45 mm syringe filters (nylon Acrodisc, Gelman Sciences, Ann Arbor, MI, USA) and analysed without further dilution. The separation of arsenic compounds by using RP-mHPLC was accomplished using a mobile phase consisting of 0.1% trifluoroacetic acid (Sigma, St. Louis, MO, USA) and 5–10% v/v methanol in water. For the RP ion-pairing (IP) mHPLC separations, tetrabutylammonium hydroxide (Alfa Products, Danvers, MA, USA) was used as the ion-pairing reagent at concentrations ranging from 1 to 5 mm.Malonic acid (Sigma) was used to adjust the mobile phase pH. The analytical mHPLC column, used for both RP and RP-IP chromatography, was a 150 mm 3 1 mm id stainless-steel column packed with Spherisorb 3 mm C18 material (Isco). HPLC mobile phases were filtered and de-gassed using a 35 mm all-glass filter holder (Millipore, Bedford, MA, USA), fitted with a 0.45 mm hydrophilic nylon filter (Cuno, Meriden, CT, USA) for aqueous solvents and a 0.5 mm PTFE filter (Cole-Parmer, Chicago, IL, USA) for organic solvents.Chromatographic standards were prepared from 1000 mg l21 As aqueous solutions of the following arsenic compounds: parsanilic acid (p-ASA) (Eastman Organic Chemicals, Rochester, NY, USA), 3-nitro-4-hydroxyphenylarsonic acid (roxarsone) (ICN Biochemicals, Cleveland, OH, USA), 4-nitrophenylarsonic acid (4-NPAA) (Aldrich, Milwaukee, WI, USA), dimethylarsinic acid (DMA) (Sigma), 4-hydroxyphenylarsonic acid (4-OH) (Eastman Kodak), disodium methylarsonate (MMA) (Chemical Service, West Chester, PA, USA), sodium meta-arsenite (Aite) (Sigma), and sodium arsenate (Aate) (Sigma).Procedures Time resolved acquisition (TRA) software provided by VG was used to acquire data during mHPLC–ICP-MS experiments. Data files of response versus time were exported as ASCII files and analysed using commercial spreadsheet software. The mFI– and mHPLC–ICP-MS response was optimised for the detection of arsenic at m/z 75 using carrier flow rates of 80 and 40 ml min21, respectively.The optimisation was carried out by employing a 60 ml loop fitted on a ceramic injector (SVI- 6U7) (Analyticon Instruments, Springfield, NJ, USA). The large volume loop provided quasi-continuous sample introduction, which lasted long enough for signal optimisation to be achieved. For the HPLC optimisation a 60 ml loop was connected post-column; this provided a signal equivalent to that obtained in the continuous flow mode.All instrument parameters were varied iteratively to reach the apparent optimum response. Results and Discussion Separation of Arsenic Animal Feed Additives Using RP- mHPLC–ICP-MS RP-mHPLC methods were developed and used for the separation of four phenylarsonic acids and two inorganic arsenic compounds. A mobile phase consisting of 0.1% trifluoroacetic acid (TFA) in water, with a variable percentage (5–15% v/v) of methanol, was used for their separation.The mHPLC stationary phase consisted of silica based C18 material (3 mm particle diameter, 12% carbon loading). The chromatograms obtained are presented in Fig. 1. Present in the injected sample was 4-hydroxyphenylarsonic acid (4-OH). Even though this compound has not been used as an animal feed additive, it may prove to be a decomposition product or metabolite of roxarsone or may adequately serve as a chromatographic internal standard. As mentioned previously, the RP-mHPLC method developed allows for the separation of inorganic arsenic species from various of the phenylarsonic acids.This separation is of particular importance, primarily because of the relatively high toxicity of the inorganic arsenic species. Thus, the RP chromatographic method can be used to monitor arsenic animal feed additives for the presence of inorganic arsenic impurities, and also to detect possible degradation of the phenylarsonic acids to their respective inorganic forms. As shown in Fig. 1, the extensive retention of the NO2-containing arsenicals (roxarsone and 4-NPAA) by the LC stationary phase results in excessive Fig. 1 RP-mHPLC–ICP-MS traces for six arsenic compounds. The mobile phase consisted of water (0.1% TFA) and 5–15% methanol, applied at a flow rate of 40 ml min21. 1064 Analyst, October 1997, Vol. 122peak tailing. A larger amount of methanol in the mobile phase reduces peak tailing significantly, but also provides poorer overall resolution.In addition, the methanol content affects the sensitivity of the ICP-MS detector. Signal enhancement for arsenic in the presence of organic solvents has been observed and reported extensively by others.19,20 This occurrence has major implications, especially when using gradient-elution methods which make use of variable amounts of organic solvents in conjunction with HPLC–ICP-MS. It has been proposed that the presence of organic solvents in the plasma alters the energetics of the ICP.Even when the introduction of organic solvent into the plasma is discontinued, a relatively long period of time is required before the ICP-MS sensitivity returns to its pre-solvent level. To explore further the possible use of RP chromatography for the speciation of phenylarsonic acids in environmental samples, we investigated the effects of naturally occurring arsenicals on the RP chromatograms. The RP-mHPLC separation of various phenylarsonic acids (arsenic animal feed additives) from naturally occurring arsenic compounds is presented in Fig. 2. These chromatograms were obtained with no methanol present in the mobile phase. Less efficient separations occurred in the presence of even small amounts of methanol ( < 0.5%). The mobile phase flow rate also had a pronounced effect on separation efficiency. The best separation was obtained at the lowest-tested flow rate (15 ml min21). The main drawback, however, was the incomplete separation of DMA from p-ASA.In addition, excessive peak broadening occurred in the low flow rate regime ( < 30 ml min21). Flow rates > 55 ml min21 were not considered, mainly because of the resulting high back-pressure ( > 3000 psi), which could potentially damage the mHPLC column. It should also be mentioned that when the mobile phase does not contain any methanol, the phenylarsonic acids roxarsone and 4-NPAA do not elute from the HPLC column. Thus, a preliminary chromatographic run using a mobile phase containing 5–15% methanol must be conducted in order to investigate the presence of the two nitro-containing arsenicals (roxarsone and 4-NPAA).Separation of Arsenic Animal Feed Additives Using RP-IP- mHPLC–ICP-MS Because RP chromatography provides poor separation of the arsenic animal feed additives from some of the naturally occurring arsenic compounds (Aite, Aate, MMA and DMA), alternative chromatographic modes of separation were investigated and further optimised to provide efficient separations.To accomplish this, RP-IP-mHPLC was further investigated. To assist in reaching the optimum conditions required for such separations, a plot of the apparent charge (Qapp) of each of the arsenic compounds as a function of mobile phase pH was constructed (Fig. 3). This plot provides a qualitative estimate of the extent of interaction of the anion-pairing reagent with the arsenic-containing analyte under various pH conditions.Qapp for arsenate was calculated as follows: Qapp AsO HAsO H AsO AsO HAsO H AsO H AsO = - + + + + + - - - - - - 3 2 4 3 4 2 2 4 4 3 4 2 2 4 3 4 [ ] [ ] [ ] [ ] [ ] [ ] [ ] The Qapp values for the other arsenic compounds were derived similarly. The pKa values21 used for these calculations were as follows: pKa[Aite] = 9.2, pKa[Aate] = 2.2, 6.98, 11.5, pKa[DMA] = 1.3,22 6.3, pKa[MMA] = 3.41, 8.18, pKa[p- ASA] = 2, 4.02, 8.92. The chromatogram presented in Fig. 4, obtained under RPIP- mHPLC conditions, exhibits sufficient separation of p-ASA Fig. 2 RP-mHPLC–ICP-MS traces for p-ASA (D) and 4-OH (E) in the presence of naturally occurring arsenic compounds: arsenite and arsenate (A), methylarsonic acid (B) and dimethylarsinic acid (C). The mobile phase consisted of water (0.1% TFA). The flow rates varied between 15 and 55 ml min21. Fig. 3 Apparent charge of arsenic compounds as a function of solution pH. Arrows indicate normal operating range of pH for reversed-phase anionpairing chromatography of arsenic acids.Fig. 4 RP-IP-mHPLC–ICP-MS traces for five arsenic compounds. The mobile phase consisted of 1 mm tetrabutylammonium hydroxide and 0.5% methanol in de-ionised water (pH 5.28), applied at a flow rate of 40 ml min21. Analyst, October 1997, Vol. 122 1065from four other arsenic compounds which have been found to occur naturally in various environmental samples. The elution order of these compounds at pH 5.3 is in agreement with that predicted by the Qapp versus pH plot (Fig. 3). A compound with high Qapp is expected to interact to a greater extent with the tetrabutylammonium ion-pairing reagent. Stronger interaction results in longer retention times. However, it should be noted that factors other than pH significantly influence the chromatographic behavior of the arsenic compounds. The most important of these factors include the ion-pairing reagent concentration and the methanol content of the mobile phase.Separation of p-ASA from Arsenic Compounds Present in Reference Materials Initially the effects of large amounts of NaCl on the chromatographic separation of arsenic compounds were investigated. The resulting chromatograms from these experiments are presented in Fig. 5. It was observed that the presence of 0.1% NaCl in the sample had a significant effect on the chromatographic separations (retention times, peak shapes), particularly when a low ion-pairing concentration (1–2 mm TBAH) was used.In order to minimise this problem, which is especially pronounced when speciating arsenic in biological materials such as urine, 5 mm TBAH was used. Also, the mobile phase pH was adjusted to allow for the separation of chloride from the other arsenic compounds, thus eliminating potential interferences caused by ArCl+. Freeze-dried urine (NIST SRM 2670) containing ‘normal levels’ of arsenic (as indicated by NIST) was analysed for its arsenic content.The freeze-dried urine sample was reconstituted with de-ionized water and then spiked with p-ASA and 4-OH. The resulting chromatogram is shown in Fig. 6. Resolution between DMA and MMA was sacrificed by appropriately adjusting the mobile phase pH in order to achieve improved separation between the Cl2 and arsenate species present. The determination of arsenate is of considerable importance mainly because its toxicity is substantially higher than that of MMA or DMA.Good separation of 4-OH, used as an internal standard in this case, was also accomplished. It should be noted that the original (non-spiked) urine SRM did not contain any measurable amounts of p-ASA or 4-OH. It should also be noted that the RP anion-pairing chromatography used in this study is not capable of differentiating between arsenobetaine and arsenite, as these two compounds co-elute. Also, there exists the possibility that other non-retained arsenic species may co-elute in this front-end peak. Arsenobetaine is normally present in human urine following the consumption of seafood.23 Previous investigations using this particular reference material have shown that indeed the first peak is the result of arsenite and arsenobetaine.18,24 Solutions referred to as Trace Elements in Water (NIST SRM 1643a) and Trace Metals in Drinking Water (High-Purity Standards) were also spiked with p-ASA and 4-OH, and subsequently analysed for their content of arsenic species (Fig. 7). The spiked arsenicals were separated from arsenite and arsenate, which were originally present in the water reference materials. Furthermore, San Joaquin Soil (NIST SRM 2709) was extracted using 0.3 m H3PO4 and then spiked with p-ASA and 4-OH. The resulting chromatogram is presented in Fig. 8. The soil extract, prior to spiking, was found to contain only arsenate as the single arsenic compound present. The limits of detection for each arsenic compound, achieved under various modes of chromatography, are summarised in Table 1. Because of the extremely high sensitivity of the ICPMS detector, the chromatographic methods described are eminently suitable for the development of methodologies for the determination of arsenic animal feed additives in real samples. These methods are particularly suited for the analysis of samples containing limited amounts of analyte.mFI–ICP-MS of Arsenic Animal Feed Additives In previous work, a mFI–ICP-MS technique was developed and applied to the determination of total arsenic.14 Of interest in this study was to investigate further any sensitivity variations observed when analysing arsenic animal feed additives by using mFI–ICP-MS.A number of reports have claimed sensitivity variations for other organoarsenic compounds determined using continuous-flow ICP-MS, particularly for arsenicals present in marine organisms.20 So far, no sensitivity data have been reported regarding arsenic animal feed additives.Fig. 9 shows the peaks obtained for the four phenylarsenical compounds discussed in this paper. Each peak corresponds to an Fig. 5 Effect of salt and ion-pair reagent concentrations on the separation of four arsenic compounds. Fig. 6 RP-IP-mHPLC–ICP-MS trace for urine (NIST SRM 2670n) spiked with p-ASA and 4-OH. The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. 1066 Analyst, October 1997, Vol. 122injection of 80 pg of As (1 ml injections of a phenylarsenical solution containing 80 ppb of As). The fact that no sensitivity variations were observed permits the use of mFI–ICP-MS for the quantification of arsenic animal feed additives regardless of which phenylarsonic species are present in the sample and without their prior conversion into a common species by employing some form of digestion. Conclusions We have demonstrated the development and successful application of a variety of chromatographic methods for the identification of arsenic animal feed additives.More specifically, RP- mHPLC was used for the separation of arsenic animal feed additives from inorganic arsenic species. This mode of chromatography, however, did not allow the complete separation of arsenic animal-feed additives from naturally occurring organoarsenic compounds. RP–IP-mHPLC was used successfully for this purpose. Furthermore, the use of mHPLC offers the advantage of operating at very low flow rates (15–40 ml min21), thus allowing for minimisation of waste and the use of small sample volumes, and also offers flow rate compatibility with other mass spectrometric detectors such as electrospray and continuous-flow fast atom bombardment mass spectrometry.The latter feature may prove to be a great benefit for the structural identification of metabolites or decomposition products for which synthetic standards are not yet available.The use of the ICP-MS detector offers additional benefits, such as high selectivity and extremely good sensitivity, for the speciation of trace levels of arsenic in samples of environmental and biological origin. The US Environmental Protection Agency (EPA), through its Office of Research and Development (ORD), funded the research described here. This paper has been subjected to the Agency’s peer review and has been approved as an EPA publication. The US Government has a non-exclusive, royaltyfree license in and to any copyright covering this paper.Mention of trade names or commercial products does not constitute Fig. 7 RP-IP-mHPLC–ICP-MS traces for A, water (NIST SRM 1643a) spiked with p-ASA and 4-OH and B, water (High-Purity Standards, Drinking Water) spiked with p-ASA and 4-OH. The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. Fig. 8 RP-IP-mHPLC–ICP-MS trace for soil extract (NIST SRM 2709 San Joaquin Soil) spiked with p-ASA and 4-OH.The mobile phase consisted of 0.5% v/v methanol and 5 mm TBAH at pH 5.8, applied at a flow rate of 40 ml min21. Table 1 Limits of detection (defined as three times the standard deviation of the background) for arsenic compounds obtained under RP- and RP-IP-m- HPLC–ICP-MS conditions RP-IPRP- mHPLC– mHPLC– ICP-MS ICP-MS pg ml21 pg ml21 As pg As As pg As Arsenite (Aite) 0.10 0.10 0.6 0.6 Arsenate (Aate) 0.10 0.10 0.4 0.4 p-Arsanilic acid (p-ASA) 0.10 0.10 0.9 0.9 4-Hydroxyphenylarsonic acid (4-OH) 0.10 0.10 0.8 0.8 3-Nitro-4-hydroxyphenylarsonic acid (roxarsone) 0.12 0.12 n.d.* n.d. 4-Nitrophenylarsonic acid (4-NPAA) 0.26 0.26 n.d. n.d. * n.d.: not determined because the compounds do not elute from column under the specified conditions. Fig. 9 mFI–ICP-MS of arsenic animal feed additives. Each peak represents the injection of 1 ml of a solution of 80 pg ml21 As. Analyst, October 1997, Vol. 122 1067endorsement or recommendation for use. This work was performed while S.A.P. held a National Research Council/ CRD-LV Research Associateship. References 1 Gilbert, F. R., Wells, G. A. H., and Gunning, R. F., Vet. Rec., 1981, 109, 158. 2 Drugs Directorate, Health and Welfare Canada, Ottawa, Canada, 1981. 3 Compendium of Medicating Ingredients Brochures, Agriculture Canada, Ottawa, 5th edn., 1984. 4 Aschbacher, P. W., and Feil, V. J., J. Agric. Food Chem., 1991, 39, 146. 5 Edmonds, M. S., and Baker, D. H., J. Anim. Sci., 1986, 63, 553. 6 Rice, D. A., McMurray, C. H., McCracken, R. M., Bryson, D. G., and Maybin, R., Vet. Rec., 1980, 106, 312. 7 Weston, R. E., Wheals, B. B., and Kensett, M. J., Analyst, 1971, 96, 601. 8 Hoodless, R. A., and Tarrant, K. R., Analyst, 1973, 98, 502. 9 Morrison, J. L., J. Agric. Food Chem., 1968, 16, 704. 10 Maruo, M., Hirayama, N., Wada, H., and Kuwamoto, T., J. Chromatogr., 1989, 466, 379. 11 Hirayama, N., and Kuwamoto, T., J. Chromatogr., 1988, 457, 415. 12 Dodd, M., PhD Thesis, University of British Columbia, Vancouver, 1988. 13 Sapp, R. E., and Davidson, S., J. AOAC Int., 1993, 76, 956. 14 Pergantis, S. A., Heithmar, E. M., and Hinners, T. A., Anal. Chem., 1995, 67, 4530. 15 Dean, J. R., Ebdon, L., Foulkes, M. E., Crews, H. M., and Massey, R. C., J. Anal. At. Spectrom., 1994, 9, 615. 16 Pergantis, S. A., Cullen, W. R., Chow, D. T., and Eigendorf, G. K., J. Chromatogr. A, 1997, 764, 211. 17 Pergantis, S. A., Cullen, W. R., and Eigendorf, G. K., Biol. Mass Spectrom., 1994, 23, 749. 18 Pergantis, S. A., Winnik, W., and Betwoski, D., J. Anal. At. Spectrom., 1997, 12, 531. 19 Allain, P., Jaunault, L., Mauras, Y., Mermet, J.-M., and Delaporte, T., Anal. Chem., 1991, 63, 1497. 20 Larsen, E. H., and St�urup, S., J. Anal. At. Spectrom., 1994, 9, 1099. 21 Dean, J. A., in Lange’s Handbook of Chemistry, McGraw-Hill, New York, 13th edn., 1985. 22 Hansen, S. H., Larsen, E. H., Pritzl, G., and Cornett, C., J. Anal. At. Spectrom., 1992, 7, 629. 23 Le, X. C., Cullen, W. R., and Reimer, K. J., Clin. Chem., 1994, 40, 617. 24 Pergantis, S. A., Momplaisir, G.-M., Heithmar, E. M., and Hinners, T. A., in Proceedings of the 44th ASMS Conference on Mass Spectrometry and Allied Topics, Portland, Oregon, May 12–16, 1996, American Society for Mass Spectrometry, East Lansing, MI, USA, p. 21. Paper 7/02691I Received April 21, 1997 Accepted July 14, 1997 1068 Analyst, October 1997, Vol. 1

ISSN:0003-2654    

DOI:10.1039/a702691i

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Determination of Trace Amounts of Vanadyl Porphyrin in Marine Mussel Tissues by High-performance Liquid Chromatography With Both Ultraviolet/Visible and Inductively Coupled Plasma Atomic Emission Spectrometric Detection Paola Rivaro* and Roberto Frache Sezione di Chimica Analitica e Ambientale, Dipartimento di Chimica e Chimica Industriale, Universit`a di Genova, via Dodecaneso, 31-16146 Genoa, Italy An HPLC method with UV/VIS and ICP-AES detection is described for the determination of vanadyl porphyrins extracted from biological samples. A detection limit of 50 ng of vanadium was obtained.The method was used to determine these compounds following their extraction from tissues of mussels treated in laboratory experiments and collected during a ‘Mussel Watch Programme’. This allowed some conclusions about vanadium speciation in marine organisms to be made. In the tissues of mussels, collected at several sites of the monitored area, which showed high vanadium concentrations, it was possible to establish the presence of this metal in the form of organometallic compounds.Keywords: Vanadyl porphyrins; speciation; liquid chromatography; inductively coupled plasma atomic emission spectrometry; hyphenated technique; biological samples; mussel watch programme Vanadium enters the marine environment through natural processes, atmospheric fallout and human activity and it is believed that in sea-water it exists primarily as orthovanadate ions.1 Oil spills can also indroduce this element into the marine environment.In petroleum, vanadium is present in the form of organometallic compounds called vanadyl porphyrins,2,3 a group of macrocyclic aromatics consisting of a porphyrin ring, to which a metal ion is bound. Etioporphyrins and deoxophylloerythroporphyrins, with a total carbon number ranging from 29 to 39, are the most abundant series found. In the past, porphyrins have been used as biomarkers in the study of the origin and formation of petroleum.4 Recently, in environmental studies, attention has been focused on their possible use in ‘fingerprinting’ polluting oil residues (tar balls), because they are much more resistant to biodegradation and weathering than the hydrocarbon fraction.3 Despite the low concentrations in sea-water, ranging from 0.3 to 3.2 mg l21, vanadium is accumulated to relatively high levels in certain marine organisms, such as ascidians and certain molluscs.5 At low concentrations it is an essential trace element in animals and plants, but it also has some toxic or inhibitory effects, as it is able to inhibit the enzymes involved in the cationic transport across the cell membranes.6 Although information exists on vanadium levels in certain marine organisms, little is known about its speciation; there is a need, therefore, for more detailed knowledge of the chemical form of this element, in order to be able to assess this metal toxin in aquatic animals fully.Marine mussels have been widely used as biomonitors of heavy metal pollution in coastal areas. From December 1994 to October 1995 a ‘Mussel Watch Programme–Regione Liguria’ was carried out to assess the quality of water of the Ligurian Sea and to identify the possible sources of metal pollution. Metal concentrations were measured in mussels (Mytilus galloprovincialis, Lam) collected in different months at eight sampling sites of dissimilar water qualities as reported in Fig. 1. Vanadium was one of the metals taken into consideration; as shown under Results and Discussion (Fig. 4) its concentrations in mussels sampled in the Genoa oil port area and close to the town of Cogoleto were statistically significantly different from the values found at all the other sampling sites. Therefore, an investigation of vanadium speciation was conducted to identify the forms in which vanadium was present in these samples, i.e., whether as inorganic or organometallic (vanadyl porphyrin) species.Several methods have been proposed to separate different classes of petro porphyrins in oil, using either high-performance liquid chromatography (HPLC)7,8 or gas chromatography (GC)9,10 to separate the different classes of compound, coupled with different detection techniques such as atomic absorption spectrometry (AAS),11 inductively coupled plasma atomic emission spectrometry (ICP-AES)8,10 and inductively coupled plasma mass spectrometry (ICP-MS).12,13 We could not find any references to the extraction and determination of vanadyl porphyrins in matrices other than oil.In fact, all the studies published on vanadium distribution in sea-water1 or in marine organisms5,6,14 refer only to the total amount of vanadium. This work describes a method for the extraction of vanadyl porphyrins from a biological matrix (mussel tissues), and the use of ICP-AES coupled to ultraviolet/visible (UV/VIS) spectrometry as detection techniques to determine the extracted vanadium-containing compounds. The results of the application of the method to the samples collected in the above-mentioned environmental monitoring programme are also presented.Fig. 1 Sampling sites of ‘Mussel Watch Programme—Regione Liguria’. Sampling site locations: 1, Lavagna; 2, Paraggi; 3, Bogliasco; 4, Genova Darsena; 5, Genova Oil Port; 6, Pr`a; 7, Vesima; and 8, Cogoleto. Analyst, October 1997, Vol. 122 (1069–1072) 1069Experimental Reagents The methanol used was of HPLC-grade; all the other chemicals were of analytical-reagent grade and were obtained from Sigma (St.Louis, MO, USA). Vanadyl porphyrin standards (V-etio-; V-octaethyl-; and V phthalo cyanylporphyrins) were provided by ‘Stazione Sperimentale per i Combustibili’, (S. Donato Milanese, Italy). Working solutions were prepared by diluting the standards with toluene and were stable for several days. The vanadium solutions were prepared by diluting a 1000 ppm stock solution (SpectrosoL-grade, BDH, Poole, Dorset, UK).De-ionized water from a Milli-Q system (Millipore, Watford, Hertfordshire, UK) was used throughout. Apparatus HPLC The HPLC instrument used was a Varian LC system 5000 equipped with a 200 ml Rheodyne (Cotati, CA, USA) injector. Vanadyl porphyrin separations were performed on a 5 mm LiChrospher RP-8 column (250 3 4.6 mm id) (Bischoff Chromatography, Leonberg, Germany), operated at room temperature.The flow rate was 0.8 ml min21 and no gradient elution devices were used. The mobile phase was methanol– water (9 + 1). The UV/VIS detector was set at the maximum absorbance wavelength (406 nm for V-etio- and V-octaethylporphyrins or 700 nm for V phthalocyanylporphyrins). ICP-AES A Jobin–Yvon 24 spectrometer (Jobin–Yvon, Longjumeau, Paris, France) was used both to determine the total vanadium in mussel tissues and as a detector for HPLC in vanadium speciation studies.In the latter case, the ICP-AES parameters were: power, 700 W; principal argon flow rate, 16 l min21; coolant, 0.2 l min21; nebulizer, 0.7 l min21. At the HPLC–ICPAES interface, a Cetac USN 5000 ultrasonic nebulizer with the thermostat set at 140 °C and the cryostat at 28 °C was used. The working wavelength for vanadium was V(II) 292.402 nm. Sample Preparation Field experiments Mussels were collected in different months (December 1994, March 1995, June 1995, July 1995, October 1995) at eight sampling sites of the ‘Mussel Watch Programme’ as reported in Fig. 1. The tissues of 20 animals, removed from the shells, were homogenized using a Turrax high-speed homogenizer and were stored at 220 °C until analysis. Laboratory experiments This part of the work was necessary to verify whether mussels were able to concentrate vanadium in their tissues as organometallic compounds, whether any degradation processes occurred and to test the procedure for extracting vanadyl porphyrins from biological samples.Mussels, of 4–6 cm in length, obtained from a marine farm located near La Spezia, were exposed to an environment containing 100 mg l 21 V as an octaethylporphyrin solution, in a static sea-water system for 6 d. The mussels were first left in poly(propylene) tanks in seasonally adjusted, artificial seawater (15 °C) prepared according to the method of La Roche et al.15 The porphyrin solution was added daily, after changing the water.Some mussels were not exposed to the treatment and they were considered as control samples. Tissues were removed from shells, homogenized and stored at 220 °C . Analytical Procedure Total vanadium A 2 g amount of homogenized tissue (wet mass) was boiled in 10 ml of 1 m nitric acid for 2 h at 70 °C in a reflux system. The acid solution was filtered through a Schleicher–Schull blueband filter (i.e., 2 mm pore size) and diluted to the final volume (20 ml) with de-ionized water.The accuracy of the procedure was checked using the standard additions method. Extraction of vanadyl porphyrins A 2 g amount of homogenized tissue (wet mass) was added to 2 g of solid Na2SO4 and left at 60 °C overnight. The extraction from the tissue was accomplished with a Soxhlet apparatus, using toluene (200 ml for each sample). Eight extraction cycles were necessary to obtain reproducible results. The toluene was collected and evaporated to dryness by using a vacuum rotary evaporator.The samples were re-dissolved in 1 ml of methanol and 200 ml aliquots were injected onto the HPLC column. The accuracy of the method was tested by analyses of sub-samples of equal mass spiked with vanadyl porphyrin standard solution. The spiking experiments were carried out by adding to the homogenized tissue a few drops of V-etio- and V-octaethylporphyrin solutions, in order to obtain additions of 2 and 4 mg l21 as V. The usual extraction procedure was then carried out.A recovery of about 80% was determined. Results and Discussion In Fig. 2 the chromatograms of a standard mixture of V-etioand V-octaethylporphyrins are shown. As can be seen, the retention times of the vanadium peaks on the ICP-AES chromatogram are slighly shifted with respect to the UV peaks, because of the connecting tube between the output of the UV/VIS detector and the plasma torch (a dead volume of about 1 ml). The UV/VIS detector was fixed at 406 nm, because the considered porphyrins had their maximum absorbance at this wavelength.The use of an ultrasonic nebulizer allowed greater flexibility in the setting-up of a chromatographic method than the classical Meinhard nebulizer. The ultrasonic nebulizer leads to a greater lowering of the organic modifier concentration reaching the plasma torch, with a positive effect on the detection limits, owing to the low noise. Fig. 2 Chromatograms of a standard solution mixture of V-etio- (peak A) and V-octaethyl (peak B) porphyrins. 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm. 1070 Analyst, October 1997, Vol. 122It should be noted that when the chromatographic method was first set up, the separations were performed using acetonitrile–water (45 + 55) as eluent. However, when the HPLC–UV/VIS detector system was coupled with the ICP-AES instrument, this organic modifier caused instability of the plasma torch, reducing its excitation properties and consequently the analytical response, while also creating a strong background signal.This drawback was overcome by using methanol, albeit in a higher percentage in order to obtain the same retention times. Under these conditions, a detection limit of 50 ng of vanadium (3s) was obtained. The detection limit was determined by injecting 200 ml aliquots of standard solutions of different concentrations onto the HPLC column. The calibration range was linear up to 1000 ng of vanadium and the relative standard deviations (n = 5) ranged from 7 to 3%.In order to verify the applicability of the extraction and determination methods to real samples, vanadyl porphyrins in tissues of mussels exposed in laboratory experiments to an octaethylporphyrin solution were determined. In Table 1, the total vanadium concentrations found in the analysed tissues (i.e., shell, total tissues, gills, digestive glands) at the end of the incubation time are reported.As can be seen, a higher vanadium concentration was found in the digestive glands. Because data on concentration factors of organometallic vanadium compounds in marine organisms were not available in the literature, these results were compared with the results of experiments in which the animals were exposed to vanadium in an inorganic form. The concentration factor for the digestive glands was higher than the values reported in the literature, whereas for the other tissues no significant differences were found.5 In Fig. 3 the chromatograms of a digestive gland extract of mussels exposed for 6 d are shown. In terms of retention time, the peaks correspond to the octaethylporphyrin present in tissues. Examination of the chromatograms revealed that all the vanadium in the samples was present as an organometallic compound, as in the ICP-AES signal, no vanadium peaks, other than this, were found. Moreover, it was established that, after 6 d of exposure, no transformation or degradation processes had occurred.The vanadium concentration calculated on the basis of peak area is in agreement with the total amount of vanadium found, taking into account the recovery value of the extraction. However, the procedure was considered suitable for the determination of organometallic vanadium species in marine organisms and was used to identify the forms in which vanadium was present in mussel samples collected during the ‘Mussel Watch Programme—Regione Liguria’.In Fig. 4 the result of the determination of total vanadium in tissues of mussels sampled in June 1995 is shown. As can be seen, the tissues of the animals collected in the Genoa Oil Port area and close to the town of Cogoleto had vanadium contents of 0.51 and 0.84 mg g 21, respectively, which were significantly higher than the concentrations found in all the other samples. Differences were tested by means of a Student’s t-test and found to be significant at P < 0.05.To these samples and to a control sample (i.e., mussels obtained from the marine farm) the determination and extraction procedures were applied for speciation purposes. The results are shown in Fig. 5. The chromatograms of both samples show that vanadium is present as vanadyl porphyrin; in terms of retention time, the peaks correspond to this compound. This result was expected for the sample collected in the oil port. This sample shows a peak which appears to consist of two different, but very closely eluting peaks; this would indicate either the presence of two different types of vanadyl porphyrin or a degradation product.On the other hand, the sample collected near the town of Cogoleto showed unequivocally the presence of vanadium as an organometallic compound in sea-water. This fact could be explained by taking into account that in this Table 1 Concentration of total vanadium (ng g21 wet mass) in tissues of mussel not exposed (control) and in mussel after 6 d exposure to water containing V-octaethylporphyrin (100 mg l21 as V).Data represent the mean ± standard deviation of three replicates ng g21 V Concentration Tissue Control Exposed factor Shell 50 ± 2 1370 ± 200 13.7 Soft parts (whole) 20 ± 2 700 ± 80 7 Gills 10 ± 1 520 ± 40 5.2 Digestive glands 60 ± 4 3790 ± 180 37.9 Fig. 3 Chromatograms of a mussel digestive gland after exposure to Voctaethylporphyrin for 6 d. 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm.Fig. 4 Concentration levels of total vanadium (mg g 21 wet mass) found in tissues of mussels collected during the ‘Mussel Watch Programme— Regione Liguria’ (June 1995 sampling). Fig. 5 Chromatograms of mussel tissues sampled in the Genova Oil Port (A) and near Cogoleto (B) compared with a control sample (C). 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm. Analyst, October 1997, Vol. 122 1071location the oil tanker Haven sank in 1992 and that oil losses are probably still occurring.Conclusion The procedures used to extract and determine organometallic vanadium compounds in biological samples were suitable for application to environmental studies. The use of a coupled detection technique, UV/VIS–ICP-AES, was helpful in giving information on vanadium speciation in tissues of marine mussels collected during a ‘Mussel Watch Programme’, allowing a more detailed knowledge of the chemical form of this element to be obtained.The presence of vanadyl porphyrins, which are characteristic compounds of petroleum, suggested that oil spills were probably responsible for the high concentration of vanadium found at some sites of the monitored area. This work was financially supported by Regione Liguria. We are also indebted to Dr. Paolo Tittarelli of the Stazione Sperimentale per i Combustibili for providing us with the vanadyl porphyrin standards. References 1 Jeandel, C., Caisso, M., and Minster, J.F., Mar. Chem., 1987, 21, 51. 2 de Waal, W. A. J., Heemstra, S., Kraak, J. C., and Jonker, R., J., Chromatographia, 1990, 30, 38. 3 Rankin, J. C., Czernuszewich, R. S., Org. Geochem., 1993, 20, 521. 4 Quirke, J. M. E., Eglinton, G., and Maxwell, J. R., J. Am. Chem. Soc., 1979, 101, 7693. 5 Miramand, P., Guary, J. C., and Fowler, S. W., Mar. Biol., 1980, 65, 281. 6 Unsal, M., Mar. Pollut. Bull., 1982, 13, 139. 7 Sundararaman, P., Anal.Chem., 1985, 57, 2204. 8 Xu, H., and Lesage, S., J. Chromatogr., 1992, 607, 139. 9 Kaur, S., Gill, J. P., Evershed, R. P., Eglinton, G., and Maxwell, J. R., J. Chromatogr., 1989, 473, 135. 10 Zeng, Y., Seeley, J. A., Dowling, T. M., and Uden, P. C., J. High Res. Chrom., 1992, 15, 669. 11 Fish, R. H., and Komlenic, J. J., Anal. Chem., 1984, 56, 584. 12 Pretorius, W. G., Foulkes, M., Ebdon, L., and Rowland, S. J., J. High Resolut. Chromatogr., 1993, 16, 157. 13 Ebdon, L., Evans, E.H., Pretorius W. G., and Rowland, S. J., J. Anal. At. Spectrom., 1994, 9, 939. 14 Miramand, P., and Bentley, D., Mar. Biol., 1992, 114, 404. 15 La Roche, G., Eisler, R., and Tarzwell, C. M., J. Water Pollut. Control Fed., 1970, 42, 1982. Paper 7/02568H Received April 15, 1997 Accepted June 23, 1997 1072 Analyst, October 1997, Vol. 122 Determination of Trace Amounts of Vanadyl Porphyrin in Marine Mussel Tissues by High-performance Liquid Chromatography With Both Ultraviolet/Visible and Inductively Coupled Plasma Atomic Emission Spectrometric Detection Paola Rivaro* and Roberto Frache Sezione di Chimica Analitica e Ambientale, Dipartimento di Chimica e Chimica Industriale, Universit`a di Genova, via Dodecaneso, 31-16146 Genoa, Italy An HPLC method with UV/VIS and ICP-AES detection is described for the determination of vanadyl porphyrins extracted from biological samples. A detection limit of 50 ng of vanadium was obtained.The method was used to determine these compounds following their extraction from tissues of mussels treated in laboratory experiments and collected during a ‘Mussel Watch Programme’. This allowed some conclusions about vanadium speciation in marine organisms to be made.In the tissues of mussels, collected at several sites of the monitored area, which showed high vanadium concentrations, it was possible to establish the presence of this metal in the form of organometallic compounds.Keywords: Vanadyl porphyrins; speciation; liquid chromatography; inductively coupled plasma atomic emission spectrometry; hyphenated technique; biological samples; mussel watch programme Vanadium enters the marine environment through natural processes, atmospheric fallout and human activity and it is believed that in sea-water it exists primarily as orthovanadate ions.1 Oil spills can also indroduce this element into the marine environment. In petroleum, vanadium is present in the form of organometallic compounds called vanadyl porphyrins,2,3 a group of macrocyclic aromatics consisting of a porphyrin ring, to which a metal ion is bound.Etioporphyrins and deoxophylloerythroporphyrins, with a total carbon number ranging from 29 to 39, are the most abundant series found. In the past, porphyrins have been used as biomarkers in the study of the origin and formation of petroleum.4 Recently, in environmental studies, attention has been focused on their possible use in ‘fingerprinting’ polluting oil residues (tar balls), because they are much more resistant to biodegradation and weathering than the hydrocarbon fraction.3 Despite the low concentrations in sea-water, ranging from 0.3 to 3.2 mg l21, vanadium is accumulated to relatively high levels in certain marine organisms, such as ascidians and certain molluscs.5 At low concentrations it is an essential trace element in animals and plants, but it also has some toxic or inhibitory effects, as it is able to inhibit the enzymes involved in the cationic transport across the cell membranes.6 Although information exists on vanadium levels in certain marine organisms, little is known about its speciation; there is a need, therefore, for more detailed knowledge of the chemical form of this element, in order to be able to assess this metal toxin in aquatic animals fully.Marine mussels have been widely used as biomonitors of heavy metal pollution in coastal areas.From December 1994 to October 1995 a ‘Mussel Watch Programme–Regione Liguria’ was carried out to assess the quality of water of the Ligurian Sea and to identify the possible sources of metal pollution. Metal concentrations were measured in mussels (Mytilus galloprovincialis, Lam) collected in different months at eight sampling sites of dissimilar water qualities as reported in Fig. 1. Vanadium was one of the metals taken into consideration; as shown under Results and Discussion (Fig. 4) its concentrations in mussels sampled in the Genoa oil port area and close to the town of Cogoleto were statistically significantly different from the values found at all the other sampling sites.Therefore, an investigation of vanadium speciation was conducted to identify the forms in which vanadium was present in these samples, i.e., whether as inorganic or organometallic (vanadyl porphyrin) species. Several methods have been proposed to separate different classes of petro porphyrins in oil, using either high-performance liquid chromatography (HPLC)7,8 or gas chromatography (GC)9,10 to separate the different classes of compound, coupled with different detection techniques such as atomic absorption spectrometry (AAS),11 inductively coupled plasma atomic emission spectrometry (ICP-AES)8,10 and inductively coupled plasma mass spectrometry (ICP-MS).12,13 We could not find any references to the extraction and determination of vanadyl porphyrins in matrices other than oil.In fact, all the studies published on vanadium distribution in sea-water1 or in marine organisms5,6,14 refer only to the total amount of vanadium. This work describes a method for the extraction of vanadyl porphyrins from a biological matrix (mussel tissues), and the use of ICP-AES coupled to ultraviolet/visible (UV/VIS) spectrometry as detection techniques to determine the extracted vanadium-containing compounds. The results of the application of the method to the samples collected in the above-mentioned environmental monitoring programme are also presented.Fig. 1 Sampling sites of ‘Mussel Watch Programme—Regione Liguria’. Sampling site locations: 1, Lavagna; 2, Paraggi; 3, Bogliasco; 4, Genova Darsena; 5, Genova Oil Port; 6, Pr`a; 7, Vesima; and 8, Cogoleto. Analyst, October 1997, Vol. 122 (1069–1072) 1069Experimental Reagents The methanol used was of HPLC-grade; all the other chemicals were of analytical-reagent grade and were obtained from Sigma (St.Louis, MO, USA). Vanadyl porphyrin standards (V-etio-; V-octaethyl-; and V phthalo cyanylporphyrins) were provided by ‘Stazione Sperimentale per i Combustibili’, (S. Donato Milanese, Italy). Working solutions were prepared by diluting the standards with toluene and were stable for several days. The vanadium solutions were prepared by diluting a 1000 ppm stock solution (SpectrosoL-grade, BDH, Poole, Dorset, UK). De-ionized water from a Milli-Q system (Millipore, Watford, Hertfordshire, UK) was used throughout.Apparatus HPLC The HPLC instrument used was a Varian LC system 5000 equipped with a 200 ml Rheodyne (Cotati, CA, USA) injector. Vanadyl porphyrin separations were performed on a 5 mm LiChrospher RP-8 column (250 3 4.6 mm id) (Bischoff Chromatography, Leonberg, Germany), operated at room temperature. The flow rate was 0.8 ml min21 and no gradient elution devices were used. The mobile phase was methanol– water (9 + 1).The UV/VIS detector was set at the maximum absorbance wavelength (406 nm for V-etio- and V-octaethylporphyrins or 700 nm for V phthalocyanylporphyrins). ICP-AES A Jobin–Yvon 24 spectrometer (Jobin–Yvon, Longjumeau, Paris, France) was used both to determine the total vanadium in mussel tissues and as a detector for HPLC in vanadium speciation studies. In the latter case, the ICP-AES parameters were: power, 700 W; principal argon flow rate, 16 l min21; coolant, 0.2 l min21; nebulizer, 0.7 l min21.At the HPLC–ICPAES interface, a Cetac USN 5000 ultrasonic nebulizer with the thermostat set at 140 °C and the cryostat at 28 °C was used. The working wavelength for vanadium was V(II) 292.402 nm. Sample Preparation Field experiments Mussels were collected in different months (December 1994, March 1995, June 1995, July 1995, October 1995) at eight sampling sites of the ‘Mussel Watch Programme’ as reported in Fig. 1. The tissues of 20 animals, removed from the shells, were homogenized using a Turrax high-speed homogenizer and were stored at 220 °C until analysis.Laboratory experiments This part of the work was necessary to verify whether mussels were able to concentrate vanadium in their tissues as organometallic compounds, whether any degradation processes occurred and to test the procedure for extracting vanadyl porphyrins from biological samples. Mussels, of 4–6 cm in length, obtained from a marine farm located near La Spezia, were exposed to an environment containing 100 mg l 21 V as an octaethylporphyrin solution, in a static sea-water system for 6 d.The mussels were first left in poly(propylene) tanks in seasonally adjusted, artificial seawater (15 °C) prepared according to the method of La Roche et al.15 The porphyrin solution was added daily, after changing the water. Some mussels were not exposed to the treatment and they were considered as control samples.Tissues were removed from shells, homogenized and stored at 220 °C . Analytical Procedure Total vanadium A 2 g amount of homogenized tissue (wet mass) was boiled in 10 ml of 1 m nitric acid for 2 h at 70 °C in a reflux system. The acid solution was filtered through a Schleicher–Schull blueband filter (i.e., 2 mm pore size) and diluted to the final volume (20 ml) with de-ionized water. The accuracy of the procedure was checked using the standard additions method. Extraction of vanadyl porphyrins A 2 g amount of homogenized tissue (wet mass) was added to 2 g of solid Na2SO4 and left at 60 °C overnight.The extraction from the tissue was accomplished with a Soxhlet apparatus, using toluene (200 ml for each sample). Eight extraction cycles were necessary to obtain reproducible results. The toluene was collected and evaporated to dryness by using a vacuum rotary evaporator. The samples were re-dissolved in 1 ml of methanol and 200 ml aliquots were injected onto the HPLC column. The accuracy of the method was tested by analyses of sub-samples of equal mass spiked with vanadyl porphyrin standard solution.The spiking experiments were carried out by adding to the homogenized tissue a few drops of V-etio- and V-octaethylporphyrin solutions, in order to obtain additions of 2 and 4 mg l21 as V. The usual extraction procedure was then carried out. A recovery of about 80% was determined. Results and Discussion In Fig. 2 the chromatograms of a standard mixture of V-etioand V-octaethylporphyrins are shown.As can be seen, the retention times of the vanadium peaks on the ICP-AES chromatogram are slighly shifted with respect to the UV peaks, because of the connecting tube between the output of the UV/VIS detector and the plasma torch (a dead volume of about 1 ml). The UV/VIS detector was fixed at 406 nm, because the considered porphyrins had their maximum absorbance at this wavelength. The use of an ultrasonic nebulizer allowed greater flexibility in the setting-up of a chromatographic method than the classical Meinhard nebulizer.The ultrasonic nebulizer leads to a greater lowering of the organic modifier concentration reaching the plasma torch, with a positive effect on the detection limits, owing to the low noise. Fig. 2 Chromatograms of a standard solution mixture of V-etio- (peak A) and V-octaethyl (peak B) porphyrins. 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm. 1070 Analyst, October 1997, Vol. 122It should be noted that when the chromatographic method was first set up, the separations were performed using acetonitrile–water (45 + 55) as eluent. However, when the HPLC–UV/VIS detector system was coupled with the ICP-AES instrument, this organic modifier caused instability of the plasma torch, reducing its excitation properties and consequently the analytical response, while also creating a strong background signal.This drawback was overcome by using methanol, albeit in a higher percentage in order to obtain the same retention times. Under these conditions, a detection limit of 50 ng of vanadium (3s) was obtained. The detection limit was determined by injecting 200 ml aliquots of standard solutions of different concentrations onto the HPLC column. The calibration range was linear up to 1000 ng of vanadium and the relative standard deviations (n = 5) ranged from 7 to 3%. In order to verify the applicability of the extraction and determination methods to real samples, vanadyl porphyrins in tissues of mussels exposed in laboratory experiments to an octaethylporphyrin solution were determined.In Table 1, the total vanadium concentrations found in the analysed tissues (i.e., shell, total tissues, gills, digestive glands) at the end of the incubation time are reported. As can be seen, a higher vanadium concentration was found in the digestive glands. Because data on concentration factors of organometallic vanadium compounds in marine organisms were not available in the literature, these results were compared with the results of experiments in which the animals were exposed to vanadium in an inorganic form.The concentration factor for the digestive glands was higher than the values reported in the literature, whereas for the other tissues no significant differences were found.5 In Fig. 3 the chromatograms of a digestive gland extract of mussels exposed for 6 d are shown.In terms of retention time, the peaks correspond to the octaethylporphyrin present in tissues. Examination of the chromatograms revealed that all the vanadium in the samples was present as an organometallic compound, as in the ICP-AES signal, no vanadium peaks, other than this, were found. Moreover, it was established that, after 6 d of exposure, no transformation or degradation processes had occurred. The vanadium concentration calculated on the basis of peak area is in agreement with the total amount of vanadium found, taking into account the recovery value of the extraction.However, the procedure was considered suitable for the determination of organometallic vanadium species in marine organisms and was used to identify the forms in which vanadium was present in mussel samples collected during the ‘Mussel Watch Programme—Regione Liguria’. In Fig. 4 the result of the determination of total vanadium in tissues of mussels sampled in June 1995 is shown.As can be seen, the tissues of the animals collected in the Genoa Oil Port area and close to the town of Cogoleto had vanadium contents of 0.51 and 0.84 mg g 21, respectively, which were significantly higher than the concentrations found in all the other samples. Differences were tested by means of a Student’s t-test and found to be significant at P < 0.05. To these samples and to a control sample (i.e., mussels obtained from the marine farm) the determination and extraction procedures were applied for speciation purposes.The results are shown in Fig. 5. The chromatograms of both samples show that vanadium is present as vanadyl porphyrin; in terms of retention time, the peaks correspond to this compound. This result was expected for the sample collected in the oil port. This sample shows a peak which appears to consist of two different, but very closely eluting peaks; this would indicate either the presence of two different types of vanadyl porphyrin or a degradation product.On the other hand, the sample collected near the town of Cogoleto showed unequivocally the presence of vanadium as an organometallic compound in sea-water. This fact could be explained by taking into account that in this Table 1 Concentration of total vanadium (ng g21 wet mass) in tissues of mussel not exposed (control) and in mussel after 6 d exposure to water containing V-octaethylporphyrin (100 mg l21 as V).Data represent the mean ± standard deviation of three replicates ng g21 V Concentration Tissue Control Exposed factor Shell 50 ± 2 1370 ± 200 13.7 Soft parts (whole) 20 ± 2 700 ± 80 7 Gills 10 ± 1 520 ± 40 5.2 Digestive glands 60 ± 4 3790 ± 180 37.9 Fig. 3 Chromatograms of a mussel digestive gland after exposure to Voctaethylporphyrin for 6 d. 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm. Fig. 4 Concentration levels of total vanadium (mg g 21 wet mass) found in tissues of mussels collected during the ‘Mussel Watch Programme— Regione Liguria’ (June 1995 sampling).Fig. 5 Chromatograms of mussel tissues sampled in the Genova Oil Port (A) and near Cogoleto (B) compared with a control sample (C). 1: UV/VIS detection at 406 nm; 2: ICP-AES signal of V(II) at 292.402 nm. Analyst, October 1997, Vol. 122 1071location the oil tanker Haven sank in 1992 and that oil losses are probably still occurring. Conclusion The procedures used to extract and determine organometallic vanadium compounds in biological samples were suitable for application to environmental studies. The use of a coupled detection technique, UV/VIS–ICP-AES, was helpful in giving information on vanadium speciation in tissues of marine mussels collected during a ‘Mussel Watch Programme’, allowing a more detailed knowledge of the chemical form of this element to be obtained.The presence of vanadyl porphyrins, which are characteristic compounds of petroleum, suggested that oil spills were probably responsible for the high concentration of vanadium found at some sites of the monitored area. This work was financially supported by Regione Liguria. We are also indebted to Dr. Paolo Tittarelli of the Stazione Sperimentale per i Combustibili for providing us with the vanadyl porphyrin standards. References 1 Jeandel, C., Caisso, M., and Minster, J. F., Mar. Chem., 1987, 21, 51. 2 de Waal, W. A. J., Heemstra, S., Kraak, J. C., and Jonker, R., J., Chromatographia, 1990, 30, 38. 3 Rankin, J. C., Czernuszewich, R. S., Org. Geochem., 1993, 20, 521. 4 Quirke, J. M. E., Eglinton, G., and Maxwell, J. R., J. Am. Chem. Soc., 1979, 101, 7693. 5 Miramand, P., Guary, J. C., and Fowler, S. W., Mar. Biol., 1980, 65, 281. 6 Unsal, M., Mar. Pollut. Bull., 1982, 13, 139. 7 Sundararaman, P., Anal. Chem., 1985, 57, 2204. 8 Xu, H., and Lesage, S., J. Chromatogr., 1992, 607, 139. 9 Kaur, S., Gill, J. P., Evershed, R. P., Eglinton, G., and Maxwell, J. R., J. Chromatogr., 1989, 473, 135. 10 Zeng, Y., Seeley, J. A., Dowling, T. M., and Uden, P. C., J. High Res. Chrom., 1992, 15, 669. 11 Fish, R. H., and Komlenic, J. J., Anal. Chem., 1984, 56, 584. 12 Pretorius, W. G., Foulkes, M., Ebdon, L., and Rowland, S. J., J. High Resolut. Chromatogr., 1993, 16, 157. 13 Ebdon, L., Evans, E. H., Pretorius W. G., and Rowland, S. J., J. Anal. At. Spectrom., 1994, 9, 939. 14 Miramand, P., and Bentley, D., Mar. Biol., 1992, 114, 404. 15 La Roche, G., Eisler, R., and Tarzwell, C. M., J. Water Pollut. Control Fed., 1970, 42, 1982. Paper 7/02568H Received April 15, 1997 Accepted June 23, 1997 1072 Analyst, October 1997, Vol. 122

ISSN:0003-2654    

DOI:10.1039/a702568h

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Supported Liquid Membrane Enrichment Combined With Atomic Absorption Spectrometry for the Determination of Lead in Urine Nii-Kotey Djanea, Ingvar A. Bergdahlb, Kuria Ndung’ua, Andrejs Sch�utzb, Gillis Johanssona and Lennart Mathiasson*a a Department of Analytical Chemistry, University of Lund, P.O. Box 124, S-221 00 Lund, Sweden b Department of Occupational and Environmental Medicine, University Hospital, S-221 85 Lund, Sweden Supported liquid membrane (SLM) methodology was used for sample clean-up and enrichment of lead in urine prior to determination by AAS.Lead ions at pH 3 were extracted across a membrane solution containing 40% m/m di-2-ethylhexylphosphoric acid dissolved in kerosene and back-extracted into an acceptor solution of 1 mol l21 nitric acid. The mechanism of mass transfer is a proton gradient across the membrane. The concentration range investigated was between 5 and 80 ng ml21 and the extraction time was varied between 0.5 and 4 h, leading to enrichment factors of up to 200.The extraction efficiency was about 95%. The detection limit, expressed as 3s of five replicate determinations, using a 45 min enrichment of a urine sample low in lead, was 0.1 and 6.0 mg l21 for ETAAS, and FAAS respectively. The results obtained by the developed method agreed with those obtained by direct ICP-MS determinations for reference urine samples and samples from occupationally lead-exposed workers. The linear correlation coefficient was 0.97, the slope of the regression line was 1.06 and the intercept was 20.37 mg l21.The 95% confidence intervals for the slope and the intercept were 0.95 to 1.18 and 23.9 to 3.1, respectively. The results at the 95% confidence level for reference urine material were of 91 ± 1.5 and 92 ± 2.0 mg l21 for ICP-MS and SLM–AAS, respectively, which agreed well with the recommended value of 90 mg l21 (range 83–97 mg l21). Keywords: Lead; urine; supported liquid membranes; atomic absorption spectrometry; inductively coupled plasma mass spectrometry; carry-over effects Different enrichment techniques have been described for the determination of trace levels of heavy metals in urine, e.g., preconcentration by electrodeposition in combination with electrothermal atomic absorption spectrometry (ETTAS)1–3 and stripping techniques.4–6 Sample preparation methods based on liquid–liquid extraction7–10 are now gradually being replaced by solid-phase extractions. In the latter case, the facilitated online connection to the final analysis equipment increases the possibilities for automated analysis.11,12 Another modern approach to sample clean-up and enrichment is to use supported liquid membrane (SLM) methodology.This technique has been shown to give efficient clean-up of organic analytes in urine and plasma samples.13 In a recent paper,14 we reported the use of di- 2-ethylhexylphosphoric acid (DEHPA) as the extractant in an SLM-based procedure for the determination of some heavy metal ions including lead in both reagent water and natural water samples.The use of a proton gradient as the main transport mechanism eliminates the use of buffers that are often contaminated with the analytes of interest. In this study, we demonstrated the possibility of extending the DEHPA extraction system to the determination of lead in a biological matrix, urine. For comparison, direct determination of lead in dilute urine samples was carried out using inductively coupled plasma mass spectrometry (ICP-MS).Experimental Equipment Atomic absorption spectrometry The ETAAS system was a Varian AA-1475, fitted with a GTA- 95 graphite tube atomiser and a programmable sample dispenser (Varian, Victoria, Australia). Deuterium lamp background correction was used. The furnace programme is given in Table 1. The flame AAS (FAAS) system was a Varian AA-6. Parameter settings, such as lamp current and wavelength, were those recommended by the manufacturer. Inductively coupled plasma The possibility of using ICP-MS for the direct determination of metals in environmental and biological samples has been demonstrated in a number of studies.15–18 For lead in blood samples, ICP-MS has proved to be very useful.19,20 The ICPMS system used was a Plasma Quad PQ2 Plus (Fison Elemental, Winsford, Cheshire, UK).SLM equipment The SLM device used was similar to one described previously, 21 except that a modification was made to facilitate membrane replacements.The device consists of two identical circular PTFE blocks (diameter 120 mm, thickness 15 mm) with grooves cut in an Archimedes spiral (depth 0.25 mm, width 1.5 mm, length 250 cm), giving channels with a total volume of 1 ml. A porous PTFE membrane with a polyethylene backing (pore size 0.2 mm, porosity 0.70, total thickness 175 mm, of which 115 mm is the polyethylene backing; Type FG, Millipore, Table 1 Graphite furnace programme: wavelength 217.0 nm, lamp current 4.0 mA, slit 1.0 nm, slit height normal, background correction on.Pyrolytic graphite-coated graphite tubes were used Argon gas Tempera- Ramp Hold flow rate/ Step ture/°C time/s time/s l min21 Drying 90 5.0 10.0 3.0 Drying 120 5.0 30.0 3.0 Ashing 400 5.0 20.0 3.0 Ashing 400 0.0 2.0 0.1 Atomization 2000 1.0 2.0 0.1 Cleaning 2200 1.0 1.0 3.0 Analyst, October 1997, Vol. 122 (1073–1077) 1073Bedford, MA, USA) was impregnated by soaking it for at least 15 min in kerosene containing 40% m/m of DEHPA. The membrane was then placed between the two PTFE blocks and clamped with a stainless-steel screw in the middle.The screw was covered with a polyetherether ketone (PEEK) tube to prevent contact between the steel surface and the solutions in the membrane channels. Chemicals All solutions were prepared from either Suprapur or analyticalreagent grade chemicals and high-purity water was obtained from a MilliQ-RO4 unit (Millipore).Polyethene bottles were used throughout and were cleaned with 4 m HNO3 for at least 2 d and rinsed with high-purity water before use. Nitric acid (Suprapur) was obtained from Merck (Darmstadt, Germany) DEHPA (95%) from Sigma Chemicals (St. Louis, MO, USA), kerosene from Kebo Lab (Malm�o, Sweden) and lead nitrate stock standard solution (1000 mg ml21) from BDH (Poole, Dorset, UK). Freeze-dried urine (batch 403125, Seronorm, Nycomed, Oslo, Norway) was used as a reference material.The recommended value of lead was 90 (range 83–97) mg l21. A synthetic urine sample was prepared by dissolving 2.08 g of creatinine (Sigma), 1.25 g of urea (BDH), 14.1 g of NaCl, 2.8 g of KCl, 0.794 g of CaCl2·2H2O, 0.88 g of MgSO4·7H2O and 1.9 ml of ammonia solution (all from Merck) in high-purity water and diluting to 1000 ml.22 Sampling and Sample Pre-treatment Urine samples were obtained from lead-exposed workers at a secondary lead smeltery.Each individual was provided with a 2 l polyethylene bottle and was instructed to collect urine at home, but not in the smeltery. All urine samples used in the preliminary experiments were obtained from occupationally non-exposed subjects. The urine samples were preserved by adjusting the pH to 3.0 with concentrated nitric acid to prevent the precipitation of calcium phosphate and the consequent loss of heavy metals due to coprecipitation or adsorption on the precipitate.Usually, urine has a pH value in the range 5.0–6.5, and 2.5–3.0 ml of concentrated nitric acid were needed for each litre of urine to adjust the pH to 3.0. The freeze-dried reference urine samples had a pH below 3.0 after reconstitution with highpurity water. Prior to SLM–AAS analysis the pH was adjusted to 3.0 with ammonia solution. The final concentration of analyte in these samples solution was 20 mg l21. These samples were processed in the same manner as the other samples for 45 min and at a flow-rate of 0.9 ml min21.For the ICP-MS determinations, the reconstituted freeze-dried urine sample was analysed after a 10-fold dilution. Enrichment and AAS Determinations Fig. 1 shows the set-up for preconcentration of lead. It is similar to that described previously.14 Briefly, the sample is pumped at a flow rate of 1 ml min21 with the peristaltic pump A. After thrichment, the acceptor solution is pumped with the peristaltic pump B to a vial placed in a fraction collector (D).The pneumatic valve A directs the sample stream to the donor channel in the SLM unit, and valve B directs fresh solution to the acceptor channel and keeps a closed loop on the stagnant acceptor during the enrichment step. Cleaning of the tube between valve C and the fraction collector D is done with peristaltic pump C for off-line analysis using ETAAS or FAAS. Normally, 2 ml of acceptor solution are used to transfer the enriched sample to a vial.To reduce possible carry-over effects due to slow kinetics at the membrane–acceptor interface, the acceptor channel was usually washed with 10 ml of the stripping solution at a flow rate of 1.0 ml min21 after each preconcentration procedure. pH changes in the acceptor during the extraction process were monitored as described previously.14 Experiments concerning analyte recovery were carried out on natural urine samples spiked with known concentrations of lead.The extent of carry-over effects was investigated by collecting and analysing 2 ml fractions of the acceptor solution, collected 0, 5 and 10 min after the preconcentration. Each collection procedure took about 160 s. ICP-MS Determinations The urine samples were acidified by adding 2% v/v concentrated nitric acid (Aristar, BDH), and left overnight. Lead was determined according to a method used previously for whole blood.19 Briefly, the samples were analysed after a 10-fold dilution with a solution containing EDTA, Triton X-100 and ammonia., The samples were introduced into the ICP-MS equipment in a flow injection-like manner (segmented flow mode; 14 s uptake, 24 s acquisition).Bismuth and thallium were used as internal standards. Validation The validation of the proposed method was carried out by comparing the results of ICP-MS and SLM–AAS determinations. Results and Discussion Influence of Enrichment Time on Acceptor pH During the use of DEHPA as carrier in the extraction of lead from natural waters, a donor pH of 3 and an acceptor pH of 0 have been found to be suitable for the mass transfer of analytes to the acceptor side.14 Accordingly, these conditions were chosen for the investigation of the influence of the enrichment and the pH of the acceptor solution.Fig. 2 shows the influence of enrichment time on acceptor pH. It is seen that the pH on the acceptor side rises steadily and reaches a value of about 1.6 after approximately 4 h.Extraction Efficiency Table 2 gives the extraction efficiencies from synthetic urine and diluted spiked natural urine. The extraction efficiencies for lead in spiked natural urine and synthetic urine are 93 and 95%, respectively, which are in general slightly higher than those obtained in diluted urine. Using a two-tailed Student’s t-test, the Fig. 1 Experimental set-up for metal enrichment. 1074 Analyst, October 1997, Vol. 122differences between diluted and undiluted urine samples was not significant at the 95% confidence level except for the samples diluted to 30 and 50%.However, the difference is not significant at the 99% confidence level. Hence there is no reason to dilute the urine samples as this will lead to higher detection limits. Further, it may be concluded that physiologically normal variations between the urine samples will not influence the extraction efficiency. For reliable quantification it is important to work under conditions where the enrichment factor is constant.The influence of the enrichment time on the extraction efficiency at the normal sample flow rate of 1 ml min21 is shown in Table 3. The extraction efficiency (Table 3) is independent of the enrichment time up to at least 2 h. The value at 4 h is lower but the difference between this value and the other values in Table 3 is not significant at the 95% confidence level using a twotailed Student’s t-test. We have found previously that the pH gradient across the membrane should be about 2 pH units in order to keep the mass transfer rate constant.14 After 4 h, the difference is about 1.4 pH units (Fig. 2). However, the driving force is the difference in H+ concentration across the membrane. Since the pH scale is logarithmic, the difference in pH may be relatively smaller at high H+ concentrations, without giving significant changes in extraction efficiency, which is clearly demonstrated here. The change in extraction efficiency with time depending on the increase in pH on the acceptor side does not seriously affect the long-term stability.Experiments have been run continuously at a flow rate of 1 ml min21 on the same membrane for periods of up to 48 h with replacement of the acceptor solution every 2.5 h without any significant changes in extraction efficiency. In this respect, the behaviour is similar to what has been observed previously for lead in reagent water and natural waters, where the long-term stability was confirmed for periods of at least 30 and 200 h for natural and reagent waters, respectively.14 The detection limit is inversely related to the enrichment time as long as the extraction efficiency is independent of time. A 4 h run at 1 ml min21 (with negligible decrease in extraction efficiency) results in a processed sample volume of 240 ml.With an acceptor volume of approximately 1.0 ml and an extraction efficiency for lead of about 90%, an enrichment factor of more than 200 is obtained.Carry-over Effects in the Membrane System In some SLM systems, the mass transfer kinetics at the membrane–acceptor interface can be slow, which has been shown previously, e.g., in the enrichment of surfactants using ion-pair formation.23 It is therefore necessary in the investigation of new SLM applications always to consider the extent of carry-over effects, which may occur as a result of limited mass transfer kinetics in the actual system.Table 4 shows the extraction efficiencies of lead at different elution times after the completion of an enrichment. The first 2 ml fraction was collected immediately after the enrichment, the second after a waiting time of 5 min, and the third after 10 min. From Table 4, it may be concluded that a 10 min wash of the acceptor channel is sufficient to remove about 98% of the enriched lead. This makes memory effects negligible, provided that the concentration difference between samples is reasonable.Quantification Effects of initial donor concentration on lead recovery The concentration dependence was tested by spiking urine samples with lead in the concentration range 5–80 mg l21 and performing a 30 min SLM enrichment at a flow rate of 1.0 ml min21 followed by analysis by ETAAS for spiked Fig. 2 Change in acceptor pH with time during SLM processing of a urine sample at pH 3.0 with a flow rate of 1 ml min21. Acceptor solution, 1.0 m nitric acid; membrane liquid, 40% m/m DEHPA in kerosene.Table 2 Extraction efficiency of lead in spiked synthetic urine and spiked natural urine of various dilutions in high-purity water (n = 5). Standard deviations are given in parentheses. Lead concentration, 50 mg l21; sample pH, 3.0; acceptor solution, 1.0 m HNO3; enrichment time, 60 min; flow rate approximately 1.0 ml min21 Urine in Extraction Urine in Extraction water (%) efficiency (%) water (%) efficiency (%) 5 91.6 (3.1) 60 90.5 (2.4) 10 88.5 (5.5) 70 88.9 (3.2) 20 89.0 (3.7) 80 94.4 (8.5) 30 88.5 (2.5) 90 93.5 (2.9) 40 90.0 (3.8) 100 93.2 (2.9) 50 89.2 (2.1) Synthetic urine 95.3 (3.0) Table 3 Extraction efficiency of lead in spiked natural urine at different enrichment times. Standard deviations are given in parentheses.Lead concentration 50 mg l21; sample, pH 3.0; acceptor solution, 1.0 m HNO3; sample flow rate, approximately 1.0 ml min21 Extraction efficiency (%) Time/h (SD) n 0.5 94.3 (1.3) 6 1.0 94.7 (1.4) 6 2.0 95.3 (1.2) 6 4.0 93.5 (4.0) 4 Table 4 Carry-over effects using SLM methodology for enriching lead in a spiked urine sample investigated using two different porous membrane filters.Two enrichments on each filter. Average values are given. Standard deviations are given in parentheses. Lead concentration, 5.5 mg l21; membrane liquid, 40% m/m DEHPA in kerosene; sample pH, 3.0; acceptor solution, 1.0 m HNO3; flow rate 0.90–1.0 ml min21; enrichment time, 100 min.The eluted volume from the acceptor after each waiting time was 2 ml Waiting time/ Extraction efficiency min (%) 0 93.1 (0.5) 93.0 (0.8) 5 5.3 (1.1) 4.3 (1.5) 10 0.4 (2.0) 0.5 (1.0) Analyst, October 1997, Vol. 122 1075concentrations below 25 mg l21 and FAAS for higher concentrations. The results are given in Table 5. Applying a Student’s t-test for the differences in Table 5 between the recoveries obtained at different concentrations and the average recovery value of 93.8% revealed that there were no significant differences at the 95% confidence level for any of the concentrations in the range considered. Detection limit The detection limit was determined as three times the standard deviation of five replicate determinations of lead in a urine sample from an occupationally unexposed individual.In two different experiments with enrichment times of 45 min, with a flow rate of 1 ml min21 and ETAAS determination, the detection limit was 0.1 mg l21.This value is well below the concentrations found in unexposed individuals. The corresponding value for FAAS is 6.0 mg l21. This is sufficient for measuring samples from workers with moderate lead exposure. The concentration of lead in occupationally exposed workers is normally above 10 mg l21. Validation In the validation using the reference urine samples, SLM–AAS gave a lead value of 92.0 ± 2.0 mg l21 (five replicates) at the 95% confidence level assuming a Student’s t distribution.This was in accordance with the recommended value of 90 (range 83–97) mg l21 (five replicates). The corresponding result for ICP-MS was 91.0 ± 1.5 mg l21 (five replicates) at the 95% confidence level. The results obtained with SLM–AAS and ICP-MS agreed well with each other and with the certified values. The plot of the results in this comparison (samples from lead-exposed workers) is illustrated in Fig. 3. Linear correlation24 based on 22 samples gave a slope of 1.06, an intercept of 20.37 mg l21 and a correlation coefficient of 0.97.The 95% confidence intervals for the slope and intercept were 0.95 to 1.18 and 23.9 to 3.1, respectively. Since these intervals include the ideal values of 1 and 0 for the slope and the intercept, respectively, there is no evidence of a systematic difference between the two sets of results. The relative deviations between the ICP-MS and the SLM– AAS values are larger for low concentrations.At low concentrations, lead was determined by ETAAS in the enriched samples. The larger deviations within this concentration range (below 20 mg l21) are probably due to technical problems related to the background correction in ETAAS. In some experiments using ETAAS, RSD values of !20% were encountered. Since FAAS generally gives good precision (RSD Å 4% in the considered concentration interval here), the final analysis could be based entirely on FAAS by prolonging the enrichment time for low-concentration samples.The longer times needed do not necessarily decrease the sample throughput. With a multi-channel system in an off-line configuration, many samples can be simultaneously enriched by the SLM technique with low labour input. The relatively low cost of FAAS equipment, low operational cost, robustness and good stability of the FAAS system together with short analysis times makes SLM–FAAS a powerful alternative. The results in Tables 2, 4 and 5 indicate that neither the matrix composition nor the initial analyte concentration influences the extraction efficiency.However, these experiments were performed on the same sample bulk. In the comparison experiments, samples were collected from different individuals. The overall agreement between the results obtained with SLM– FAAS and ICP-MS in this comparison indicates that individual variations in the urine composition do not influence the results in any significant way.Furthermore, the present sample cleanup process using SLM does not seem to affect the results in any significant way. Using SLM with a cation-exchange extractant, negatively charged lead complexes, for example, could have been excluded from the membrane transport process. Even species which under the given conditions in the donor solution can form neutral complexes with lead might not contribute to the mass transfer of lead across the membrane, thereby decreasing the extraction efficiency to values below 100%.The most important compounds to be considered are probably chloride, phosphate, sulfate, citrate and oxalate ions, which are normally present in undiluted urine at concentrations of about 6000, 1200, 180 and 10–50 mg l21, respectively. Also, high concentrations of urea and creatinine could have some effects. However, these changes must be small. In the validation experiment the creatinine concentration varied between 5.2 and 20.9 mm.According to the literature,25 the normal variation of creatinine in urine is 0.13–0.22 mm kg21. Hence for a person with a mass of 70 kg, the normal level of creatinine should vary from 9.1 to 15.4 mm. Obviously the variation of creatinine in the samples investigated exceeded this range without leading to a significant influence on the extraction efficiency of lead. Concerning urea, the dilution experiments in Table 3 show that dilution has a negligible effect on the recovery of lead.The range was further investigated by comparing the recovery of lead in synthetic urine samples with normal urea concentrations (20 mm) with samples with about three times higher (62 mm) concentrations. It was found that the difference between the average values (based on three determinations at 20 mm and six determinations at 62 mm) were less than 1%. Hence the method developed seems to be well suited for clinical measurements. Table 5 Recovery of lead in natural urine spiked with different lead concentrations.The background concentration of lead, 0.9 mg l21, obtained as an average for six unspiked natural urine samples, was subtracted before calculating the extraction efficiency. The number of experiments at each concentration was six. Standard deviations are given in parentheses. Lead concentration, 5–80 mg l21; sample pH, 3.0; acceptor solution, 1.0 m HNO3; enrichment time, 30 min; flow rate, 1.0 ml min21 Lead concentration/ Recovery mg l21 (%) 5 92.4 (3.5) 10 94.8 (5.9) 20 94.0 (5.8) 50 95.3 (1.7) 80 92.7 (6.5) Fig. 3 Validation of the SLM–AAS methodology for lead determination by comparison with results obtained with ICP-MS. Sample pH, 3.0; acceptor solution, 1.0 m nitric acid; membrane liquid, 40% m/m DEHPA in kerosene. 1076 Analyst, October 1997, Vol. 122Conclusion We have shown that SLM combined with AAS can be used for the determination of lead in urine samples. By extending the enrichment time to 2 h, a detection limit of 2 mg l21 is obtainable using FAAS, thus covering the whole range of interest at occupational exposure.The methodology could be extended to other metals such as Cd, Cu and Cr, which will also give high extraction efficiencies in the chosen SLM system. For ICP-MS determinations, the flexibility of the SLM technique may be potentially useful when sample clean-up procedures are necessary, to avoid spectral interferences from certain elements.This work was made possible by financial support from the Swedish Environmental Protection Agency and the Medical Faculty at Lund University. The authors are grateful to Docent J. Å. J�onsson for fruitful suggestions. They also thank workers of Boliden Bergs�oe AB, Landskrona, for providing the urine samples. Mr. A. Ekholm and Mr. F. Malcus are acknowledged for skilful technical assistance. References 1 Zhang, G., Li, J., Fu, D., Hao, D., and Xiang, P., Talanta, 1993, 40, 409. 2 Boxing, X., Ming, X. T., Neng, S. M., and Ahi, F. Y., Talanta, 1985, 32, 1016. 3 Wolff, E. W., Landy, M. P., and Pell, D. A., Anal. Chem., 1981, 53, 1566. 4 Connor, M., Dempsey, E., Smyth, M. R., and Richardson, D. H. S., Electroanalysis, 1991, 3, 331. 5 Agraz, R., Seilla, M. T., Pinilla, J. M., and Hernandez, L., Electroanalysis, 1991 393. 6 Agraz, R., Sevilla, M. T., and Hernandez, L., Anal. Chim. Acta, 1993, 273, 205. 7 Bruland, K. W., Franks, R. P., Knauer, G. A., and Martin, J.H., Anal. Chim. Acta, 1979, 105, 233. 8 Pedersen, B., Willems, M., and Jørgensen, S., Analyst, 1980, 105, 119. 9 Smith, C. L., Motooka, J. M., and Willson, W. R., Anal. Lett., 1984, 17, 1715. 10 Ping, L., Matsumoto, K., and Fuwa, K., Anal. Chim. Acta, 1983, 147, 205. 11 Sperling, M., Yin, X., and Weiz, B., J. Anal. At. Spectrom., 1991, 6, 615. 12 Malamas, F., Bengtsson, M., and Johansson, G., Anal. Chim. Acta, 1984, 160, 1. 13 Lindegråd, B., J�onsson, J.-Å., and Mathiasson, L., J.Chromatogr., 1992, 573, 191. 14 Djane, N.-K., Ndung’u, K., Malcus, F., Johansson, G., and Mathiasson, L., Fresenius’ J. Anal. Chem., 1997, 358, 822. 15 Lu, P.-L., Huang, K.-S., and Jiang, S.-J., Anal. Chim. Acta, 1993, 284, 181. 16 Houk, R. S., Fassel, V. A., Svec, H. J., Gray, A. L., and Taylor, C. E., Anal. Chem., 1980, 52, 2283. 17 Douglas, D. J., and Houk, R. S., Prog. Anal. At. Spectrom., 1985, 8, 1. 18 Vanhoe, H., J. Trace Elem. Electrolytes Health Dis., 1993, 7, 131. 19 Bergdahl, I. A., Schutz, A., Geraldsson, L., Jensen, A., and Skerfving, S., Scand. J. Work Environ. Health, in the press. 20 Sch�utz, A., Bergdahl, I. A., Ekholm, A., and Skerfving, S., J. Occup. Med., 1996, 53, 736. 22 Papantoni, M., Djane, N.-K., Ndung’u, K., J�onsson, J.-Å., and Mathiasson, L., Analyst, 1995, 120, 1471. 22 Krushevska, A., Barnes, R. M., and Amarasiriwaradena, C., Analyst, 1993, 118, 1175. 23 Miliotis, T., Knutsson, M., J�onsson, J. Å., and Mathiasson, L., Int.J. Environ. Anal. Chem., 1996, 64, 35. 24 Miller, J. C., and Miller, J. N., Statistics for Analytical Chemistry, Ellis Horwood, Chichester, 3rd edn., 1994. 25 The Merck Manual of Diagnosis and Therapy, ed. Berkow, R., and Fletcher, J. A., Merck, Rahway, NJ, 16th edn., 1992. Paper 7/02340E Received April 7, 1997 Accepted June 23, 1997 Analyst, October 1997, Vol. 122 1077 Supported Liquid Membrane Enrichment Combined With Atomic Absorption Spectrometry for the Determination of Lead in Urine Nii-Kotey Djanea, Ingvar A.Bergdahlb, Kuria Ndung’ua, Andrejs Sch�utzb, Gillis Johanssona and Lennart Mathiasson*a a Department of Analytical Chemistry, University of Lund, P.O. Box 124, S-221 00 Lund, Sweden b Department of Occupational and Environmental Medicine, University Hospital, S-221 85 Lund, Sweden Supported liquid membrane (SLM) methodology was used for sample clean-up and enrichment of lead in urine prior to determination by AAS.Lead ions at pH 3 were extracted across a membrane solution containing 40% m/m di-2-ethylhexylphosphoric acid dissolved in kerosene and back-extracted into an acceptor solution of 1 mol l21 nitric acid. The mechanism of mass transfer is a proton gradient across the membrane. The concentration range investigated was between 5 and 80 ng ml21 and the extraction time was varied between 0.5 and 4 h, leading to enrichment factors of up to 200. The extraction efficiency was about 95%.The detection limit, expressed as 3s of five replicate determinations, using a 45 min enrichment of a urine sample low in lead, was 0.1 and 6.0 mg l21 for ETAAS, and FAAS respectively. The results obtained by the developed method agreed with those obtained by direct ICP-MS determinations for reference urine samples and samples from occupationally lead-exposed workers. The linear correlation coefficient was 0.97, the slope of the regression line was 1.06 and the intercept was 20.37 mg l21.The 95% confidence intervals for the slope and the intercept were 0.95 to 1.18 and 23.9 to 3.1, respectively. The results at the 95% confidence level for reference urine material were of 91 ± 1.5 and 92 ± 2.0 mg l21 for ICP-MS and SLM–AAS, respectively, which agreed well with the recommended value of 90 mg l21 (range 83–97 mg l21). Keywords: Lead; urine; supported liquid membranes; atomic absorption spectrometry; inductively coupled plasma mass spectrometry; carry-over effects Different enrichment techniques have been described for the determination of trace levels of heavy metals in urine, e.g., preconcentration by electrodeposition in combination with electrothermal atomic absorption spectrometry (ETTAS)1–3 and stripping techniques.4–6 Sample preparation methods based on liquid–liquid extraction7–10 are now gradually being replaced by solid-phase extractions.In the latter case, the facilitated online connection to the final analysis equipment increases the possibilities for automated analysis.11,12 Another modern approach to sample clean-up and enrichment is to use supported liquid membrane (SLM) methodology.This technique has been shown to give efficient clean-up of organic analytes in urine and plasma samples.13 In a recent paper,14 we reported the use of di- 2-ethylhexylphosphoric acid (DEHPA) as the extractant in an SLM-based procedure for the determination of some heavy metal ions including lead in both reagent water and natural water samples. The use of a proton gradient as the main transport mechanism eliminates the use of buffers that are often contaminated with the analytes of interest.In this study, we demonstrated the possibility of extending the DEHPA extraction system to the determination of lead in a biological matrix, urine. For comparison, direct determination of lead in dilute urine samples was carried out using inductively coupled plasma mass spectrometry (ICP-MS).Experimental Equipment Atomic absorption spectrometry The ETAAS system was a Varian AA-1475, fitted with a GTA- 95 graphite tube atomiser and a programmable sample dispenser (Varian, Victoria, Australia). Deuterium lamp background correction was used. The furnace programme is given in Table 1. The flame AAS (FAAS) system was a Varian AA-6. Parameter settings, such as lamp current and wavelength, were those recommended by the manufacturer. Inductively coupled plasma The possibility of using ICP-MS for the direct determination of metals in environmental and biological samples has been demonstrated in a number of studies.15–18 For lead in blood samples, ICP-MS has proved to be very useful.19,20 The ICPMS system used was a Plasma Quad PQ2 Plus (Fison Elemental, Winsford, Cheshire, UK).SLM equipment The SLM device used was similar to one described previously, 21 except that a modification was made to facilitate membrane replacements. The device consists of two identical circular PTFE blocks (diameter 120 mm, thickness 15 mm) with grooves cut in an Archimedes spiral (depth 0.25 mm, width 1.5 mm, length 250 cm), giving channels with a total volume of 1 ml.A porous PTFE membrane with a polyethylene backing (pore size 0.2 mm, porosity 0.70, total thickness 175 mm, of which 115 mm is the polyethylene backing; Type FG, Millipore, Table 1 Graphite furnace programme: wavelength 217.0 nm, lamp current 4.0 mA, slit 1.0 nm, slit height normal, background correction on.Pyrolytic graphite-coated graphite tubes were used Argon gas Tempera- Ramp Hold flow rate/ Step ture/°C time/s time/s l min21 Drying 90 5.0 10.0 3.0 Drying 120 5.0 30.0 3.0 Ashing 400 5.0 20.0 3.0 Ashing 400 0.0 2.0 0.1 Atomization 2000 1.0 2.0 0.1 Cleaning 2200 1.0 1.0 3.0 Analyst, October 1997, Vol. 122 (1073–1077) 1073Bedford, MA, USA) was impregnated by soaking it for at least 15 min in kerosene containing 40% m/m of DEHPA.The membrane was then placed between the two PTFE blocks and clamped with a stainless-steel screw in the middle. The screw was covered with a polyetherether ketone (PEEK) tube to prevent contact between the steel surface and the solutions in the membrane channels. Chemicals All solutions were prepared from either Suprapur or analyticalreagent grade chemicals and high-purity water was obtained from a MilliQ-RO4 unit (Millipore). Polyethene bottles were used throughout and were cleaned with 4 m HNO3 for at least 2 d and rinsed with high-purity water before use.Nitric acid (Suprapur) was obtained from Merck (Darmstadt, Germany) DEHPA (95%) from Sigma Chemicals (St. Louis, MO, USA), kerosene from Kebo Lab (Malm�nd lead nitrate stock standard solution (1000 mg ml21) from BDH (Poole, Dorset, UK). Freeze-dried urine (batch 403125, Seronorm, Nycomed, Oslo, Norway) was used as a reference material. The recommended value of lead was 90 (range 83–97) mg l21.A synthetic urine sample was prepared by dissolving 2.08 g of creatinine (Sigma), 1.25 g of urea (BDH), 14.1 g of NaCl, 2.8 g of KCl, 0.794 g of CaCl2·2H2O, 0.88 g of MgSO4·7H2O and 1.9 ml of ammonia solution (all from Merck) in high-purity water and diluting to 1000 ml.22 Sampling and Sample Pre-treatment Urine samples were obtained from lead-exposed workers at a secondary lead smeltery. Each individual was provided with a 2 l polyethylene bottle and was instructed to collect urine at home, but not in the smeltery.All urine samples used in the preliminary experiments were obtained from occupationally non-exposed subjects. The urine samples were preserved by adjusting the pH to 3.0 with concentrated nitric acid to prevent the precipitation of calcium phosphate and the consequent loss of heavy metals due to coprecipitation or adsorption on the precipitate. Usually, urine has a pH value in the range 5.0–6.5, and 2.5–3.0 ml of concentrated nitric acid were needed for each litre of urine to adjust the pH to 3.0.The freeze-dried reference urine samples had a pH below 3.0 after reconstitution with highpurity water. Prior to SLM–AAS analysis the pH was adjusted to 3.0 with ammonia solution. The final concentration of analyte in these samples solution was 20 mg l21. These samples were processed in the same manner as the other samples for 45 min and at a flow-rate of 0.9 ml min21. For the ICP-MS determinations, the reconstituted freeze-dried urine sample was analysed after a 10-fold dilution.Enrichment and AAS Determinations Fig. 1 shows the set-up for preconcentration of lead. It is similar to that described previously.14 Briefly, the sample is pumped at a flow rate of 1 ml min21 with the peristaltic pump A. After the enrichment, the acceptor solution is pumped with the peristaltic pump B to a vial placed in a fraction collector (D). The pneumatic valve A directs the sample stream to the donor channel in the SLM unit, and valve B directs fresh solution to the acceptor channel and keeps a closed loop on the stagnant acceptor during the enrichment step.Cleaning of the tube between valve C and the fraction collector D is done with peristaltic pump C for off-line analysis using ETAAS or FAAS. Normally, 2 ml of acceptor solution are used to transfer the enriched sample to a vial. To reduce possible carry-over effects due to slow kinetics at the membrane–acceptor interface, the acceptor channel was usually washed with 10 ml of the stripping solution at a flow rate of 1.0 ml min21 after each preconcentration procedure.pH changes in the acceptor during the extraction process were monitored as described previously.14 Experiments concerning analyte recovery were carried out on natural urine samples spiked with known concentrations of lead. The extent of carry-over effects was investigated by collecting and analysing 2 ml fractions of the acceptor solution, collected 0, 5 and 10 min after the preconcentration.Each collection procedure took about 160 s. ICP-MS Determinations The urine samples were acidified by adding 2% v/v concentrated nitric acid (Aristar, BDH), and left overnight. Lead was determined according to a method used previously for whole blood.19 Briefly, the samples were analysed after a 10-fold dilution with a solution containing EDTA, Triton X-100 and ammonia., The samples were introduced into the ICP-MS equipment in a flow injection-like manner (segmented flow mode; 14 s uptake, 24 s acquisition).Bismuth and thallium were used as internal standards. Validation The validation of the proposed method was carried out by comparing the results of ICP-MS and SLM–AAS determinations. Results and Discussion Influence of Enrichment Time on Acceptor pH During the use of DEHPA as carrier in the extraction of lead from natural waters, a donor pH of 3 and an acceptor pH of 0 have been found to be suitable for the mass transfer of analytes to the acceptor side.14 Accordingly, these conditions were chosen for the investigation of the influence of the enrichment and the pH of the acceptor solution.Fig. 2 shows the influence of enrichment time on acceptor pH. It is seen that the pH on the acceptor side rises steadily and reaches a value of about 1.6 after approximately 4 h. Extraction Efficiency Table 2 gives the extraction efficiencies from synthetic urine and diluted spiked natural urine.The extraction efficiencies for lead in spiked natural urine and synthetic urine are 93 and 95%, respectively, which are in general slightly higher than those obtained in diluted urine. Using a two-tailed Student’s t-test, the Fig. 1 Experimental set-up for metal enrichment. 1074 Analyst, October 1997, Vol. 122differences between diluted and undiluted urine samples was not significant at the 95% confidence level except for the samples diluted to 30 and 50%.However, the difference is not significant at the 99% confidence level. Hence there is no reason to dilute the urine samples as this will lead to higher detection limits. Further, it may be concluded that physiologically normal variations between the urine samples will not influence the extraction efficiency. For reliable quantification it is important to work under conditions where the enrichment factor is constant. The influence of the enrichment time on the extraction efficiency at the normal sample flow rate of 1 ml min21 is shown in Table 3.The extraction efficiency (Table 3) is independent of the enrichment time up to at least 2 h. The value at 4 h is lower but the difference between this value and the other values in Table 3 is not significant at the 95% confidence level using a twotailed Student’s t-test. We have found previously that the pH gradient across the membrane should be about 2 pH units in order to keep the mass transfer rate constant.14 After 4 h, the difference is about 1.4 pH units (Fig. 2). However, the driving force is the difference in H+ concentration across the membrane. Since the pH scale is logarithmic, the difference in pH may be relatively smaller at high H+ concentrations, without giving significant changes in extraction efficiency, which is clearly demonstrated here. The change in extraction efficiency with time depending on the increase in pH on the acceptor side does not seriously affect the long-term stability.Experiments have been run continuously at a flow rate of 1 ml min21 on the same membrane for periods of up to 48 h with replacement of the acceptor solution every 2.5 h without any significant changes in extraction efficiency. In this respect, the behaviour is similar to what has been observed previously for lead in reagent water and natural waters, where the long-term stability was confirmed for periods of at least 30 and 200 h for natural and reagent waters, respectively.14 The detection limit is inversely related to the enrichment time as long as the extraction efficiency is independent of time.A 4 h run at 1 ml min21 (with negligible decrease in extraction efficiency) results in a processed sample volume of 240 ml. With an acceptor volume of approximately 1.0 ml and an extraction efficiency for lead of about 90%, an enrichment factor of more than 200 is obtained. Carry-over Effects in the Membrane System In some SLM systems, the mass transfer kinetics at the membrane–acceptor interface can be slow, which has been shown previously, e.g., in the enrichment of surfactants using ion-pair formation.23 It is therefore necessary in the investigation of new SLM applications always to consider the extent of carry-over effects, which may occur as a result of limited mass transfer kinetics in the actual system.Table 4 shows the extraction efficiencies of lead at different elution times after the completion of an enrichment. The first 2 ml fraction was collected immediately after the enrichment, the second after a waiting time of 5 min, and the third after 10 min.From Table 4, it may be concluded that a 10 min wash of the acceptor channel is sufficient to remove about 98% of the enriched lead. This makes memory effects negligible, provided that the concentration difference between samples is reasonable. Quantification Effects of initial donor concentration on lead recovery The concentration dependence was tested by spiking urine samples with lead in the concentration range 5–80 mg l21 and performing a 30 min SLM enrichment at a flow rate of 1.0 ml min21 followed by analysis by ETAAS for spiked Fig. 2 Change in acceptor pH with time during SLM processing of a urine sample at pH 3.0 with a flow rate of 1 ml min21. Acceptor solution, 1.0 m nitric acid; membrane liquid, 40% m/m DEHPA in kerosene. Table 2 Extraction efficiency of lead in spiked synthetic urine and spiked natural urine of various dilutions in high-purity water (n = 5).Standard deviations are given in parentheses. Lead concentration, 50 mg l21; sample pH, 3.0; acceptor solution, 1.0 m HNO3; enrichment time, 60 min; flow rate approximately 1.0 ml min21 Urine in Extraction Urine in Extraction water (%) efficiency (%) water (%) efficiency (%) 5 91.6 (3.1) 60 90.5 (2.4) 10 88.5 (5.5) 70 88.9 (3.2) 20 89.0 (3.7) 80 94.4 (8.5) 30 88.5 (2.5) 90 93.5 (2.9) 40 90.0 (3.8) 100 93.2 (2.9) 50 89.2 (2.1) Synthetic urine 95.3 (3.0) Table 3 Extraction efficiency of lead in spiked natural urine at different enrichment times.Standard deviations are given in parentheses. Lead concentration 50 mg l21; sample, pH 3.0; acceptor solution, 1.0 m HNO3; sample flow rate, approximately 1.0 ml min21 Extraction efficiency (%) Time/h (SD) n 0.5 94.3 (1.3) 6 1.0 94.7 (1.4) 6 2.0 95.3 (1.2) 6 4.0 93.5 (4.0) 4 Table 4 Carry-over effects using SLM methodology for enriching lead in a spiked urine sample investigated using two different porous membrane filters.Two enrichments on each filter. Average values are given. Standard deviations are given in parentheses. Lead concentration, 5.5 mg l21; membrane liquid, 40% m/m DEHPA in kerosene; sample pH, 3.0; acceptor solution, 1.0 m HNO3; flow rate 0.90–1.0 ml min21; enrichment time, 100 min. The eluted volume from the acceptor after each waiting time was 2 ml Waiting time/ Extraction efficiency min (%) 0 93.1 (0.5) 93.0 (0.8) 5 5.3 (1.1) 4.3 (1.5) 10 0.4 (2.0) 0.5 (1.0) Analyst, October 1997, Vol. 122 1075concentrations below 25 mg l21 and FAAS for higher concentrations. The results are given in Table 5.Applying a Student’s t-test for the differences in Table 5 between the recoveries obtained at different concentrations and the average recovery value of 93.8% revealed that there were no significant differences at the 95% confidence level for any of the concentrations in the range considered.Detection limit The detection limit was determined as three times the standard deviation of five replicate determinations of lead in a urine sample from an occupationally unexposed individual. In two different experiments with enrichment times of 45 min, with a flow rate of 1 ml min21 and ETAAS determination, the detection limit was 0.1 mg l21. This value is well below the concentrations found in unexposed individuals.The corresponding value for FAAS is 6.0 mg l21. This is sufficient for measuring samples from workers with moderate lead exposure. The concentration of lead in occupationally exposed workers is normally above 10 mg l21. Validation In the validation using the reference urine samples, SLM–AAS gave a lead value of 92.0 ± 2.0 mg l21 (five replicates) at the 95% confidence level assuming a Student’s t distribution. This was in accordance with the recommended value of 90 (range 83–97) mg l21 (five replicates).The corresponding result for ICP-MS was 91.0 ± 1.5 mg l21 (five replicates) at the 95% confidence level. The results obtained with SLM–AAS and ICP-MS agreed well with each other and with the certified values. The plot of the results in this comparison (samples from lead-exposed workers) is illustrated in Fig. 3. Linear correlation24 based on 22 samples gave a slope of 1.06, an intercept of 20.37 mg l21 and a correlation coefficient of 0.97.The 95% confidence intervals for the slope and intercept were 0.95 to 1.18 and 23.9 to 3.1, respectively. Since these intervals include the ideal values of 1 and 0 for the slope and the intercept, respectively, there is no evidence of a systematic difference between the two sets of results. The relative deviations between the ICP-MS and the SLM– AAS values are larger for low concentrations. At low concentrations, lead was determined by ETAAS in the enriched samples.The larger deviations within this concentration range (below 20 mg l21) are probably due to technical problems related to the background correction in ETAAS. In some experiments using ETAAS, RSD values of !20% were encountered. Since FAAS generally gives good precision (RSD Å 4% in the considered concentration interval here), the final analysis could be based entirely on FAAS by prolonging the enrichment time for low-concentration samples. The longer times needed do not necessarily decrease the sample throughput.With a multi-channel system in an off-line configuration, many samples can be simultaneously enriched by the SLM technique with low labour input. The relatively low cost of FAAS equipment, low operational cost, robustness and good stability of the FAAS system together with short analysis times makes SLM–FAAS a powerful alternative. The results in Tables 2, 4 and 5 indicate that neither the matrix composition nor the initial analyte concentration influences the extraction efficiency.However, these experiments were performed on the same sample bulk. In the comparison experiments, samples were collected from different individuals. The overall agreement between the results obtained with SLM– FAAS and ICP-MS in this comparison indicates that individual variations in the urine composition do not influence the results in any significant way. Furthermore, the present sample cleanup process using SLM does not seem to affect the results in any significant way. Using SLM with a cation-exchange extractant, negatively charged lead complexes, for example, could have been excluded from the membrane transport process. Even species which under the given conditions in the donor solution can form neutral complexes with lead might not contribute to the mass transfer of lead across the membrane, thereby decreasing the extraction efficiency to values below 100%.The most important compounds to be considered are probably chloride, phosphate, sulfate, citrate and oxalate ions, which are normally present in undiluted urine at concentrations of about 6000, 1200, 180 and 10–50 mg l21, respectively. Also, high concentrations of urea and creatinine could have some effects.However, these changes must be small. In the validation experiment the creatinine concentration varied between 5.2 and 20.9 mm. According to the literature,25 the normal variation of creatinine in urine is 0.13–0.22 mm kg21. Hence for a person with a mass of 70 kg, the normal level of creatinine should vary from 9.1 to 15.4 mm.Obviously the variation of creatinine in the samples investigated exceeded this range without leading to a significant influence on the extraction efficiency of lead. Concerning urea, the dilution experiments in Table 3 show that dilution has a negligible effect on the recovery of lead. The range was further investigated by comparing the recovery of lead in synthetic urine samples with normal urea concentrations (20 mm) with samples with about three times higher (62 mm) concentrations.It was found that the difference between the average values (based on three determinations at 20 mm and six determinations at 62 mm) were less than 1%. Hence the method developed seems to be well suited for clinical measurements. Table 5 Recovery of lead in natural urine spiked with different lead concentrations. The background concentration of lead, 0.9 mg l21, obtained as an average for six unspiked natural urine samples, was subtracted before calculating the extraction efficiency.The number of experiments at each concentration was six. Standard deviations are given in parentheses. Lead concentration, 5–80 mg l21; sample pH, 3.0; acceptor solution, 1.0 m HNO3; enrichment time, 30 min; flow rate, 1.0 ml min21 Lead concentration/ Recovery mg l21 (%) 5 92.4 (3.5) 10 94.8 (5.9) 20 94.0 (5.8) 50 95.3 (1.7) 80 92.7 (6.5) Fig. 3 Validation of the SLM–AAS methodology for lead determination by comparison with results obtained with ICP-MS. Sample pH, 3.0; acceptor solution, 1.0 m nitric acid; membrane liquid, 40% m/m DEHPA in kerosene. 1076 Analyst, October 1997, Vol. 122Conclusion We have shown that SLM combined with AAS can be used for the determination of lead in urine samples. By extending the enrichment time to 2 h, a detection limit of 2 mg l21 is obtainable using FAAS, thus covering the whole range of interest at occupational exposure.The methodology could be extended to other metals such as Cd, Cu and Cr, which will also give high extraction efficiencies in the chosen SLM system. For ICP-MS determinations, the flexibility of the SLM technique may be potentially useful when sample clean-up procedures are necessary, to avoid spectral interferences from certain elements. This work was made possible by financial support from the Swedish Environmental Protection Agency and the Medical Faculty at Lund University. The authors are grateful to Docent J. Å. J�onsson for fruitful suggestions. They also thank workers of Boliden Bergs�oe AB, Landskrona, for providing the urine samples. Mr. A. Ekholm and Mr. F. Malcus are acknowledged for skilful technical assistance. References 1 Zhang, G., Li, J., Fu, D., Hao, D., and Xiang, P., Talanta, 1993, 40, 409. 2 Boxing, X., Ming, X. T., Neng, S. M., and Ahi, F. Y., Talanta, 1985, 32, 1016. 3 Wolff, E. W., Landy, M. P., and Pell, D. A., Anal. Chem., 1981, 53, 1566. 4 Connor, M., Dempsey, E., Smyth, M. R., and Richardson, D. H. S., Electroanalysis, 1991, 3, 331. 5 Agraz, R., Seilla, M. T., Pinilla, J. M., and Hernandez, L., Electroanalysis, 1991, 3, 393. 6 Agraz, R., Sevilla, M. T., and Hernandez, L., Anal. Chim. Acta, 1993, 273, 205. 7 Bruland, K. W., Franks, R. P., Knauer, G. A., and Martin, J. H., Anal. Chim. Acta, 1979, 105, 233. 8 Pedersen, B., Willems, M., and Jørgensen, S., Analyst, 1980, 105, 119. 9 Smith, C. L., Motooka, J. M., and Willson, W. R., Anal. Lett., 1984, 17, 1715. 10 Ping, L., Matsumoto, K., and Fuwa, K., Anal. Chim. Acta, 1983, 147, 205. 11 Sperling, M., Yin, X., and Weiz, B., J. Anal. At. Spectrom., 1991, 6, 615. 12 Malamas, F., Bengtsson, M., and Johansson, G., Anal. Chim. Acta, 1984, 160, 1. 13 Lindegråd, B., J�onsson, J.-Å., and Mathiasson, L., J. Chromatogr., 1992, 573, 191. 14 Djane, N.-K., Ndung’u, K., Malcus, F., Johansson, G., and Mathiasson, L., Fresenius’ J. Anal. Chem., 1997, 358, 822. 15 Lu, P.-L., Huang, K.-S., and Jiang, S.-J., Anal. Chim. Acta, 1993, 284, 181. 16 Houk, R. S., Fassel, V. A., Svec, H. J., Gray, A. L., and Taylor, C. E., Anal. Chem., 1980, 52, 2283. 17 Douglas, D. J., and Houk, R. S., Prog. Anal. At. Spectrom., 1985, 8, 1. 18 Vanhoe, H., J. Trace Elem. Electrolytes Health Dis., 1993, 7, 131. 19 Bergdahl, I. A., Schutz, A., Geraldsson, L., Jensen, A., and Skerfving, S., Scand. J. Work Environ. Health, in the press. 20 Sch�utz, A., Bergdahl, I. A., Ekholm, A., and Skerfving, S., J. Occup. Med., 1996, 53, 736. 22 Papantoni, M., Djane, N.-K., Ndung’u, K., J�onsson, J.-Å., and Mathiasson, L., Analyst, 1995, 120, 1471. 22 Krushevska, A., Barnes, R. M., and Amarasiriwaradena, C., Analyst, 1993, 118, 1175. 23 Miliotis, T., Knutsson, M., J�onsson, J. Å., and Mathiasson, L., Int. J. Environ. Anal. Chem., 1996, 64, 35. 24 Miller, J. C., and Miller, J. N., Statistics for Analytical Chemistry, Ellis Horwood, Chichester, 3rd edn., 1994. 25 The Merck Manual of Diagnosis and Therapy, ed. Berkow, R., and Fletcher, J. A., Merck, Rahway, NJ, 16th edn., 1992. Paper 7/02340E Received April 7, 1997 Accepted June 23, 1997 Analyst, October 1997, V

ISSN:0003-2654    

DOI:10.1039/a702340e

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Effect of Sample Volume on Quantitative Analysis by Solid-phase Microextraction Part 1. Theoretical Considerations Tadeusz G�orecki† and Janusz Pawliszyn* Department of Chemistry and Waterloo Centre for Groundwater Research, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1 This paper discusses the effect of sample volume on the amount of analyte extracted from a sample by solid-phase microextraction (SPME) in two-phase (sample–fiber coating) and three-phase (sample–headspace–fiber coating) systems.Up-to-date knowledge is summarized, and new concepts are introduced. The effect of sample volume on quantification and precision of results can be neglected only in rare cases. The minimum sample volume which ensures that the amount extracted, n, is lower than 1% of the initial amount of the analyte present in the sample, as well as the volume for which exactly half of the initial amount of the analyte is extracted, have been calculated for both two- and three-phase systems.It is critical that the volumes of samples and standards are the same during analysis by SPME. Extraction kinetics in headspace analysis is dependent on the headspace capacity. If it is sufficiently large, the analyte is extracted almost exclusively from the gaseous phase, and equilibration can be very fast. On the other hand, this causes a significant loss of sensitivity. The effect of sample volume on the determination of the value of the partition coefficient, K, is also discussed. If the change in concentration of the analyte in the sample at equilibrium is not taken into account, erroneous results are obtained.Even when a proper procedure is used, there are practical limitations to the accuracy of the K value determination. Large sample volumes should always be used for K value determination, as they enable broader ranges of K values to be covered with good accuracy. Keywords: Solid-phase microextraction; sample volume; partition coefficient Solid-phase microextraction (SPME) was introduced in 1990.1 Since then, the interest in this method continues to grow very rapidly.SPME utilizes a small, fused-silica fiber coated with a polymeric stationary phase for analyte extraction from the matrix. The fiber is mounted for protection in a syringe-like device. The stationary phase can be a very viscous liquid (e.g., polydimethylsiloxane, PDMS) or a porous solid. For liquid phases, analytes are absorbed until an equilibrium is reached in the system.The amount extracted under these conditions is dependent on the partition coefficient between the sample and the coating. Sampling in SPME can be carried out directly from gaseous or liquid samples, or from the headspace over liquid or solid samples.2 SPME has been used for many applications, including the determination of substituted benzene compounds, 3–5 caffeine in beverages,6 volatile organic compounds in water,7–9 polyaromatic hydrocarbons and polychlorinated biphenyls,10 chlorinated hydrocarbons,11 phenols,12–14 pesticides, 15–22 and fatty acids,23 as well as lead and tetraethyllead.24 Although, initially, SPME was used in conjunction with GC only, its applicability has been broadened to LC25,26 and supercritical fluid chromatography (SFC).27 Theoretical aspects of SPME analysis have been studied in detail.Louch et al.28 developed the theory for two-phase systems (sample and fiber coating), and Zhang and Pawliszyn2 for three-phase systems (sample–headspace–coating). Practical aspects have been discussed in numerous papers, including those of Arthur et al.29 and Motlagh and Pawliszyn.30 Although the concepts behind SPME are relatively simple, analysis of the contents of some papers, as well as conversations with numerous users, indicate that certain aspects of SPME remain unclear or are misunderstood by many people, which can easily lead to confusion and frustration with the technique.A notorious example is the often large discrepancies between the partition coefficient (distribution constant) values reported for the same coatings and compounds by different groups, or even different researchers in the same group. These discrepancies have led Langenfeld et al.31 to propose the term ‘effective distribution constant’ to describe the partition coefficients observed for given sample–coating systems. The aim of this series of papers is to discuss several aspects of SPME analysis that might be directly responsible for the problems observed.In particular, the effect of sample volume on the amount of analyte extracted in two- and three-phase systems, as well as on determination of partition coefficients, K, will be discussed. Part 1 briefly summarizes the current knowledge and introduces several new concepts. Part 2 will present experimental verification of the ideas presented. Sample Volume versus Amount Extracted in Two-phase Systems In two-phase systems (gaseous sample–coating or liquid sample–coating) at equilibrium, the initial amount of the analyte present in the sample is distributed between the sample and the SPME fiber coating.The mass balance in such systems can be described in the following way:28 C0Vs = CHs Vs + CHf Vf (1) where C0 is the initial concentration of the analyte in the sample, Vs the sample volume, CHs the concentration of the analyte in the sample at equilibrium, CHf the concentration of the analyte in the fiber coating at equilibrium and Vf the volume of the coating.It should be emphasized that for liquid samples the above mass balance applies only to systems with no headspace, or for analytes whose Henry’s constants are so low that their amount in the sample headspace can be neglected. Partitioning between the sample and fiber coating is governed by the partition coefficient, K, also called the distribution constant: K C C = ¥ ¥ f s (2) † On leave from the Faculty of Chemistry, Technical University of Gda�nsk, Poland.Analyst, October 1997, Vol. 122 (1079–1086) 1079Combination of eqns. (1) and (2) and a few simple rearrangements yield the final expression describing the amount extracted by the SPME fiber at equilibrium, n:28 n KC V V V KV = + 0 s f s f (3) It is a common notion that the term KVf in the denominator of eqn. (3) is usually so small that it can be neglected, in which case the amount of analyte extracted by the fiber coating at equilibrium would be independent of sample volume and simply described by n = KC0Vf.While this may be true for analytes characterized by very low coating–sample partition coefficients or for samples of very large volumes, in most cases the above assumption is incorrect, which can lead to significant errors, as will be discussed in the following sections. The concentration of the analyte in the sample at equilibrium can be determined from a simple mass balance: C C V n V s s s ¥ = - 0 (4) Dividing both sides of eqn.(4) by C0 yields: C C C V n C V n C V KV V KV V V KV s s s s f s f s s f ¥ = - = - = - + = + 0 0 0 0 1 1 (5) Also, by definition C C n n s ¥ = 0 0 (6) where n is the amount of analyte extracted from a sample of volume Vs and n0 is the amount that would be extracted by the fiber from a sample of infinite volume (in which case no significant decrease in the concentration of the analyte in the sample would be observed at equilibrium).Fig. 1 presents the dependence of n/n0 on Vs for (a) a 100 mm fiber (Vf = 0.65 ml) and (b) a 30 mm fiber (Vf = 0.14 ml), for sample volumes of up to 1 l and K values ranging from 1000 to 1 000 000. This situation would be characteristic of air sampling from typical glass bulbs, and hydrocarbons from the gasoline range. According to Martos et al.,32 K Å 1000 corresponds to C7, while K Å 1 000 000 corresponds to C15.It is obvious from Fig. 1 that except for compounds with the lowest K values, sample volume has a very significant effect on the amount extracted by the fiber. Practical consequences of this are very important. Calibration of the fiber is usually performed by exposing it to a standard gaseous mixture generated in a glass bulb of known volume by the static me.33 The results of such calibrations are valid only for gaseous samples of exactly the same volume. If the fiber is exposed to ambient air (an unlimited supply of sample), the amount extracted by the fiber can be significantly larger compared with limited volume samples.On the other hand, if the containers used for grab sampling of air are smaller than the bulb used for calibration, the amount extracted will be smaller than expected from the calibration graph. In both cases, erroneous quantitative results will be obtained. It is obvious, therefore, that the same volume should be used for both the sample and the standard.Alternatively, a different calibration procedure can be used. Dynamic generation of standard gas mixtures provides unlimited volumes of gas standards, and thus can be used to calibrate the fiber for direct ambient air sampling. Alternatively, calibration based on retention parameters as described by Martos and Pawliszyn34 can be used for PDMS fibres. In this procedure no standards are necessary. Direct analysis of aqueous samples is usually performed by exposing the fiber to a small volume of sample contained in a vial.Fig. 2 presents similar dependences determined for sample volumes of up to 40 ml, and three coating thicknesses, viz., 100, 30 and 7 mm (Vf = 0.028 ml). It should be emphasized again that no headspace can be present in the vial if the two-phase system model is to be applied. The thin coating (7 mm) fibers are often used for direct sampling of semi-volatile compounds, since thicker coatings cause equilibration times to be excessively long.It is clear from Fig. 2 that the sample volume is an Fig. 1 Dependence of n/n0 on Vs for (a) 100 mm fiber (Vf = 0.65 ml), and (b) 30 mm fiber (Vf = 0.14 ml), for direct sampling from large volume samples (two-phase system). Fig. 2 Dependence of n/n0 on Vs for (a) 100 mm fiber (Vf = 0.65 ml), (b) 30 mm fiber (Vf = 0.14 ml), and (c) 7 mm fiber (Vf = 0.028 ml), for direct sampling from small volume samples (two-phase system). 1080 Analyst, October 1997, Vol. 122important parameter even for compounds with relatively small K values, especially when very small sample volumes (1 ml) are used. Moreover, headspace sampling is normaly used for volatile compounds with small K values and direct sampling is used for semi-volatile compounds with large K values. Hence, sample volume cannot usually be neglected even when the 7 mm coating is used. The course of the curves indicates that in some cases (steep part of the curve, e.g., K = 1000 for a 100 mm coating, Vs between 1 and 5 ml) small differences in sample volume can result in relatively large differences in the amount extracted, which adversely affects the precision of the determination.Care should be taken, therefore, to ensure that the sample volume during direct sampling is always the same. In view of the above discussion, it is interesting to establish what the sample volume should be in order for the amount extracted by the fiber to be insignificant compared with the amount remaining in the sample after extraction. Eqn.(3) can be used for this purpose. Let us assume that the amount extracted, n = CfVf, fulfils the above condition when it is smaller than, or equal to, 1% of the initial analyte amount in the sample, i.e., CHf Vf @ 0.01 C0Vs (7) The equation can be rearranged to yield: C C V V KV V f f s f s ¥ @ 0 0 01 0 01 @ @ . . (8) since, according to the initial assumption, C0 � CHs . Under the above conditions, eqn.(3) can be simplifid to n = KC0Vf (9) Simple rearrangement of eqn. (8) yields the final condition: VS ! 100KVf (10) For a 100 mm fiber, whose volume is approximately 0.65 ml, eqn. (10) can be written as: VS ! 0.065 K [ml] (11) It follows from eqn. (11) that even for analytes whose partition coefficient values are as low as 100, the minimum sample volume when using a 100 mm thick coating should be 6.5 ml if the effect of sample volume on the amount extracted is to be neglected.Similar reasoning can be applied to calculate the sample volume from which exactly 50% of the analyte is extracted, i.e., n = CfVf = 0.5 C0Vs. In this case 0 5 0 0 . C V KC V V V KV s s f s f = + (12) After a few rearrangements, the final condition is obtained: Vs 50% = KVf (13) or, for a 100 mm fiber, Vs = K 3 0.65 3 1023 [ml] (14) It follows from eqn. (14) that when 1 ml samples are analyzed with 100 mm fibers, 50% of the initial amount of the analyte will be extracted when the K value is approximately 1540.Typical partition coefficients of semi-volatile compounds are usually higher; it is obvious, therefore, that sample volume has a very significant effect on the amount extracted when small sample volumes are used. Sample Volume versus Amount Extracted in Three-phase Systems In most cases liquid samples are placed in vials with some headspace remaining inside. At equilibrium, by definition the chemical potentials of the analyte in all three phases (liquid sample–headspace–fiber coating) must be the same.2 A very important consequence of this fact is that the amount of the analyte extracted by the fiber at equilibrium in a three-phase system is the same independently of where the fiber is located, be it the headspace or the liquid.Consequently, in systems where a headspace is present, a different dependence should be used to calculate the amount of analyte extracted by the fiber regardless of where the fiber is located, viz:2 n KC V V KV K V V = + + 0 s f f hs hs s (15) where Khs is the headspace–liquid partition coefficient Khs = CH hs/CHs (16) (CH hs is the concentration of the analyte in the headspace at equilibrium), and Vhs is the headspace volume.Compared with a two-phase system described by eqn. (3), the difference is the additional term KhsVhs in the denominator of eqn. (4). For volatile compounds, Khs is usually close to 1, which means that headspace volume can be neglected only when it is close to zero (a two-phase system). Semi-volatile compounds have much lower values of Khs; therefore, the KhsVhs term may be negligibly small; however, such an assumption should always be verified. For obvious reasons the dependence of the amount of analyte extracted by the fiber on sample volume is much more complex in three-phase systems.However, it can be still dealt with relatively easily if the total volume of the system (sample plus headspace) remains constant.In practice, this will be the usual case, as the vial volume remains constant, and the headspace volume is the difference between the total volume and the sample volume. In such systems the concentration of an analyte in the sample at equilibrium is determined by the following mass balance: C C V C V n V s s hs hs s ¥ ¥ = - - 0 (17) Dividing both sides of eqn. (17) by C0 yields: C C C V C V n C V s hs hs s s ¥ ¥ = - - 0 0 0 1 (18) From the definition of Khs [eqn.(16)], CH hs can be replaced by KhsCHs , while n is described by eqn. (15), which yields C C K C V C V KV KV K V V s ks s hs s f f hs hs s ¥ ¥ = - - + + 0 0 1 (19) Simple rearrangements yield the final dependence: C C V KV K V V s s f hs hs s ¥ = + + 0 (20) Similarly to a two-phase system, Cs H/C0 = n/n0 when the headspace volume is constant. Fig. 3 illustrates the effect of headspace volume on the amount of analyte extracted in a system of constant volume (4, 15 and 40 ml—typical vial sizes) by a 100 mm fiber, typically used for the analysis of volatile compounds in sample headspace.To allow for better comparison of the results, the xaxis has been defined as the ratio of headspace volume to sample volume. The following values of partition coefficients have been used to calculate the course of the curves: K = 400, 2500 and 1000, and Khs = 0.15, 0.7 and 1.24, respectively. These values correspond approximately to those for chloroform, 1,1,1-trichloroethane and carbon tetrachloride.It is interesting that n/n0 is always the largest for the dependence corresponding Analyst, October 1997, Vol. 122 1081to chloroform, which has the lowest Khs value, even though its K value is also the lowest. This is because, with such a combination of Khs and K, only araction of the analyte is present in the sample headspace; therefore, the equilibrium concentration of the analyte in the sample remains relatively high.The course of the two other curves is also interesting. For small headspace volumes, n/n0 is higher for the curve corresponding to carbon tetrachloride than for that corresponding to 1,1,1-trichloroethane, which is consistent with the values of their partition coefficients, K. However, as carbon tetrachloride has the largest Khs, n/n0 drops faster for this compound than for 1,1,1-trichloroethane, as a result of which the two lines cross each other.This illustrates the significance of headspace volume on the analytical results. It is also interesting that the largest relative increase in n/n0 ( Å 44%) when moving from small to large vials is observed for 1,1,1-trichloroethane, with the highest K value. For compounds with lower K values the relative increase in the amount extracted is much lower ( Å 16 and Å 12% for chloroform and carbon tetrachloride, respectively). It is, therefore, the combination of K and Khs for a given compound that determines the magnitude of the effect of the sample volume on the amount extracted in three-phase systems with headspace.Headspace volume can be the critical factor determining the precision of the results in three-phase systems. It is relatively easy to measure sample volume accurately. However, vials are not manufactured to have exactly the same volume. Wall thicknesses and bottom shapes may differ from vial to vial. Also, the shape of the septum in a closed vial can vary from concave to convex.All these factors will affect the total volume of the system, and, therefore, the headspace volume, which is the difference between the total volume and sample volume. As illustrated in Fig. 3, for compounds with large Khs, changing the headspace volume can significantly affect the amount extracted, especially in the range of headspace volumes usually applied (Vhs/Vs @ 1). Such differences (especially related to septum shape) are usually more pronounced in small vials; therefore, worse precision can usually be expected when using such vials for headspace sampling. Similarly to two-phase systems, it is possible to calculate the minimum sample volume necessary for the amount extracted by the fiber to be insignificant compared with the amount remaining in the sample after extraction.It is again assumed that the condition is fulfilled when less than 1% of the initial amount present in the sample is extracted by the fiber, i.e., n = CHf Vf @ 0.01 C0Vs.However, in this case it cannot be assumed that CHs � Co, because even if the amount extracted by the fiber constitutes only 1% of the initial amount of the analyte, a significant fraction of the analyte might be present in the headspace. Eqn. (15) will, therefore, be used to calculate the criterion: 001 0 0 . C V KC V V KV K V V s s f f hs hs s ! + + (21) The headspace volume, Vhs, can be expressed as a fraction of the sample volume, Vhs = aVs.Simple rearrangements yield the final criterion: V KV aK s f hs ! 99 1+ (22) Compared with the criterion for a two-phase system [eqn. (10)], there is an additional term in the denominator of eqn. (22), viz., aKhs. For non-zero headspace volumes this term is always greater than 0, which means that the sample volume fulfilling the criterion is smaller in three-phase systems than in two-phase systems. This is because, ultimately, the amount of analyte extracted by the fiber is determined by the concentration of the analyte in the liquid sample at equilibrium.This concentration will be lower in three-phase systems than in two-phase systems, since a significant fraction of the analyte may be present in the headspace of the sample. As a consequence, the sensitivity of headspace SPME can only be lower than, or equal to, that of direct SPME if the system is allowed to reach equilibrium. The higher sensitivity for headspace SPME reported sometimes (especially for compounds with relatively large K and Khs values) is clearly due to non-equilibrium conditions when sampling directly from liquid samples.This is not unusual in the light of the fact that equilibration times for direct sampling from water can be as long as a few hours (especially with inefficient agitation), and the increases in the amount of analyte extracted might not be noticeable if the experiment is not carried out for sufficiently long times.In addition, during prolonged sampling, analyte losses via adsorption onto the sample vial walls, absorption by the exposed parts of the silicone rubber septum (the Teflon lining of the septum has to be pierced to introduce the fiber; since the needle of the SPME device is blunt, the lining usually breaks open exposing a small area of silicone substrate), microbial decomposition, etc., can more than outweigh the expected increase in the amount of analyte extracted, as a result of which a smaller extracted amount can actually be observed after a longer time.This and other factors, including elimination of matrix effects related to the presence of high-boiling compounds in the sample and significantly faster equilibration times, usually make headspace SPME the preferred method for the analysis of mostly volatile compounds. It is also easy to determine the sample volume from which 50% of the analyte will be extracted by the SPME fiber placed in the sample headspace.The procedure is similar to that Fig. 3 Effect of headspace volume on the amount of analyte extracted in a system of a constant volume of (a) 4, (b) 15 and (c) 40 ml by a 100 mm fiber for chloroform, 1,1,1-trichloroethane and carbon tetrachloride (see text for the values of partition coefficients used for calculations). 1082 Analyst, October 1997, Vol. 122presented for two-phase systems. The final criterion in this case is: V KV aK s f hs 50 1 % = + (23) The difference between eqns.(23) and (13) is again the additional term in the denominator, aKhs. The higher the headspace–sample partition coefficient, Khs, or the headspaceto- sample volume ratio, a, the smaller the sample volume from which 50% of the initial amount of the analyte is extracted by the SPME fiber. Effect of Headspace Capacity on Extraction Kinetics Headspace volume can have a significant effect on equilibration times (extraction kinetics). If the headspace capacity is low (small Khs), and K is large, the equilibration process is very slow.A rapid initial increase in the amount of analyte extracted is usually observed, followed by a much slower increase that lasts for a long time. The first stage corresponds to analyte extraction from the gaseous phase only. As soon as the headspace concentration of the analyte falls below the equilibrium level with respect to the aqueous phase, analyte molecules start to move from the liquid sample to the headspace.At any given moment there can only be so many molecules in the headspace, depending on the Khs value, which causes the equilibration process to be very slow. The headspace acts in this case as a bottleneck for analyte transport to the fiber. On the other hand, if the amount of the analyte extracted by the fiber at equilibrium is negligible compared with the amount present in the headspace equilibrated with the sample, only a very small amount of the analyte has actually to be transported from the liquid sample through the headspace to the fiber coating, i.e., the analyte is extracted almost exclusively from the gaseous phase, and the process is much faster than for the case described above.Assuming this situation occurs when 95% of the analyte extracted by the fiber at equilibrium comes exclusively from the headspace, the criterion that must be fulfilled can be described by eqn. (24). The assumption is reasonable, as a 5% difference usually falls within the limits of experimental error for trace SPME–GC analysis: R KV K V KV K aV = = f hs hs f hs s @ 0.05 (24) The above criterion means that the capacity of the headspace (KhsVhs) needs to be at least 20 times larger than the capacity of the fiber (KVf) to achieve rapid extraction.For a given sample volume, Vs, this c be achieved by using a sufficiently large headspace volume (corresponding to a large headspace-tosample volume ratio, a), or by increasing Khs.The latter can be accomplished by increasing the temperature or by salting the analyte out of the liquid phase. When the criterion described by eqn. (24) is fulfilled, equilibration can take as little as a few minutes, and is almost independent of the agitation conditions (provided that the analyte is equilibrated between the liquid phase and its headspace before the extraction begins). It should be emphasized, however, that great care must be exercised when moving the method from one vial size to another, as illustrated in Fig. 4. When the K value is large and/or Khs is small [Fig. 4(a)] a significant portion of the analyte extracted by the fiber has to be transported from the liquid sample through the headspace independently of the headspace volume, as evidenced by the values of R larger than 0.05 for all three values of a illustrated except for a = 2 and sample volumes larger than 28 ml (hence headspace volumes larger than 56 ml).The equilibration times will be relatively long for all vial sizes up to 40 ml. On the other hand, when the K value is smaller and/or Khs is larger, the R value can be lower than 0.05 for sample and headspace volumes often found in practice. Whether the criterion is fulfilled or not depends on the actual headspace volume [Fig. 4(b)]. For the same headspace-to-sample volume ratio, a, the criterion is fulfilled for large headspace volumes, but not for small ones. For example, when using a headspace-tosample volume ratio a = 0.5, the criterion is fulfilled for headspace volumes greater than 7 ml ( = sample volumes greater than 14 ml); therefore, under such conditions equilibration is very fast. However, when a 2 ml sample is used and the headspace volume is 1 ml, R is equal to 0.345, which indicates that a significant portion of the analyte has to be transported to the fiber from the liquid sample.As a consequence, the equilibration time will be longer even though the overall sample volume is smaller. If this is not accounted for and extraction times determined for large vials are used with small vials, the accuracy and precision of the determination suffers.It should be remembered, therefore, that the equilibration time has to be examined whenever the volume of the sample and/or headspace, or the extraction conditions (temperature, salt addition) are changed. It should also be emphasized that the increase in headspace capacity causes a significant loss of sensitivity.On the other hand, increasing the headspace capacity (which can be accomplished, for example, by increasing the extraction temperature) might dramatically shorten the equilibration times for some semi-volatile analytes while maintaining sufficient sensitivity. Both these effects need to be taken into account when developing a method involving headspace extraction. Effect of Sample Volume on the Determination of K in Two-phase Systems The fiber coating–sample partition coefficient [eqn.(2)] can be determined in several ways. The best way is to use eqn. (3). After a few rearrangements, the following dependence is obtained: K nV V C V n = - s f s ( ) 0 (25) Fig. 4 Dependence of R = KVf/KhsaVs on sample volume for three headspace-to-sample volume ratios, a = 0.5, 1 and 2, and a 100 mm PDMS fiber (Vf = 0.65 ml); (a) K = 1000, Khs = 0.25; (b) K = 250, Khs = 0.5. Analyst, October 1997, Vol. 122 1083If the amount extracted by the fiber, n, is negligible compared with the initial amount of the analyte in the sample, C0Vs, this equation is simplified to: K n C V = 0 f (26) which is a simple rearrangement of eqn. (9). Often, K is calculated directly from its definition [eqn. (2)]. In this case, if n is negligible compared with the initial amount of the analyte in the sample, C0Vs, then CHs can be substituted by C0, and the calculation is straightforward. However, it is our belief that often this substitution is made even when it is not legitimate. The consequences of this can be very significant, as will be illustrated in the following paragraphs.In both cases, the fiber volume necessary to calculate the concentration of the analyte in the fiber coating at equilibrium must be known with sufficiently good accuracy to obtain valid results. Let us introduce a new term, K apparent (Kap), this being the ratio of the concentration of the analyte in the fiber at equilibrium to the initial concentration C0: K C C ap f = ¥ 0 (27) Kap is what is calculated when depletion of the initial concentration of the analyte in the sample is not taken into account (i.e., when C0 is used instead of CHs ).The ratio of K to Kap is equal to: K K C C ap s = ¥ 0 (28) Therefore, K C C K ap s = � ¥ 0 (29) Since (5), s s s f C C V V KV ¥ = + 0 K KV V KV ap s s f = + (30) The true value of K can be calculated using the following equation derived from eqn.(29): K K V K V V = - - ap s ap f s (31) However, in many cases it will not be possible to determine K accurately by this method owing to the phenomena illustrated in Fig. 5. Fig. 5 presents the relationship between Kap and K for a wide range of partition coefficients, for two different coatings (100 and 7 mm) and three sample volumes (2, 10 and 30 ml). The dependences level off in all cases. It is obvious that the relationship reaches a certain limit, above which the same value of Kap is determined independently of the true value of K.The limit can be easily determined mathematically: lim lim K K K KV V KV V V ®¥ ®¥ = + æ è ç ö ø ÷ = ap s s f s f (32) It follows from eqn. (31) that Kap is determined only by the phase ratio Vs/Vf. The smaller the sample size (or the thicker the coating), the lower the value of Kap. It is possible that this phenomenon is behind certain very low K values reported in the literature for compounds that should have very large partition coefficients (e.g., polyaromatic hydrocarbons), especially when small sample volumes are used.Also, it is clear that when Kap is calculated instead of K, the values for thinner fiber coatings are much higher than those for thicker coatings (the difference in volume between a 100 mm fiber coating and a 7 mm coating is more than an order of magnitude). Such apparent discrepancies have also been reported in the literature.Even if proper methodology is used, owing to practical limitations it is not always possible to determine K accurately, as will be illustrated in the following paragraphs. Table 1 presents the amount of analyte extracted from a 2 ml sample and the corresponding per cent. increase in the amount extracted when going from a smaller K to a larger K for a 100 mm thick fiber coating and 1 mg l21 initial concentration, for different K values. For small K values the increase in the amount extracted when K increases is significant, although not linear as would be the case for infinite sample volumes.For large K values the increase in the amount extracted when K increases becomes insignificant and falls within the limits of experimental error. Table 2 illustrates the significance of this phenomenon. It presents the ranges of K values that are obtained when the amount of analyte extracted by the fiber, n, determined experimentally falls within ±5% (relative) of the true value, for two sample volumes (2 and 35 ml) and two fiber coating thicknesses (100 and 7 mm).A ±5% error can be assumed typical for trace analysis by SPME–GC. Fig. 5 Relationship between Kap and K for two different coatings (100 and 7 mm) and three sample volumes (2, 10 and 30 ml). Table 1 Amount of analyte extracted from a 2 ml sample and % increase in the amount extracted for a 100 mm thick fiber coating and 1 mg l21 initial concentration, for different K values K n/ng Increase (%) 100 0.0629540 — 1 000 0.4905660 717 10 000 1.5294118 212 100 000 1.9402985 28 1 000 000 1.9938650 2.8 10 000 000 1.9993848 0.28 100 000 000 1.9999385 0.3 1084 Analyst, October 1997, Vol. 122It is clear from Table 2 that practical limitations to K determinationan be very severe, especially when thick fiber coatings and low sample volumes are used. For a 2 ml sample and a 100 mm thick fiber, the error of K determination for K values around 10 000 is around ±20% for a ±5% relative error of n determination.If the true K values are higher by an order of magnitude, the values determined experimentally can range from Å 36 000 to infinity, which is of no use. In practice, owing to experimental errors, for large K values the value of n found experimentally can be larger than the initial amount of analyte in the sample. In such cases, negative K values would be obtained, which, of course, have no physical meaning. It is necessary in such cases to assume that the amount extracted, n, is equal to the initial amount of analyte in the sample (quantitative extraction), in which case infinite K values are obtained. On the other hand, K values of Å 1 000 000 can be determined with reasonable accuracy when large sample volumes (35 ml) and thin fiber coatings (7 mm) are used.It is our belief that the above-described phenomena account for most of the discrepancies between K values reported in the literature for the same substances and coatings.Other possible sources of errors include limited solubility of some analytes in water, as well as losses of analytes due to volatilization (when headspace is present in the system), absorption by exposed parts of silicone rubber septa, adsorption on glass, or biodegradation. The first two phenomena are more probable for volatile analytes, while the remaining phenomena are more significant for semi-volatile compounds, since when K values are very high, equilibration times become very long.From a practical point of view, partition coefficients, K, should always be determined using large sample volumes (nothing can be gained by using small volumes). For semi-volatile compounds characterized by large K values, only thin coatings (7 mm for PDMS) are practical for the experimental determination of K. Above a certain level, the accurate determination of K becomes virtually impossible. Conclusions Sample volume plays an extremely important role in analysis by SPME.In order to avoid errors or poor precision, care should be taken to ensure that the volumes of samples and standard solutions used for calibration are the same. In headspace SPME, the combination of K and Khs determines the magnitude of the effect of sample volume on the amount extracted by the fiber, n. With the same headspace-to-sample volume ratio, the magnitude of the effect is greatest for compounds with low Khs. The minimum sample volume which ensures that n is lower than 1% of the initial amount of analyte present in the sample (which means that the change in concentration of the analyte in the sample at equilibrium is negligible) can easily be calculated for both two- and three-phase systems, and so is the volume for which exactly half of the initial analyte amount is extracted. Variable vial volumes might cause poorer precision in headspace sampling, as only the volume of the liquid sample can be accurately controlled.Extraction kinetics in headspace sampling is determined by the headspace capacity, KhsVhs. If it is sufficiently large compared with the fiber capacity, KVf, then the analyte is extracted almost exclusively from the headspace, and equilibration can be very fast. The headspace capacity is determined by the actual headspace volume rather than by the headspace-tosample volume ratio a; therefore, dramatically different equilibration times might be observed when scaling down the method from large to small vials, even though a remains constant.On the other hand, a large headspace capacity causes a significant loss of method sensitivity. Sample volume should always be taken into account when determining K values. If the change in concentration of the analyte in the sample at equilibrium is not taken into account, erroneous results are obtained. Even when a proper procedure is used, there are practical limitations to the accuracy of the K value determination.Large sample volumes should always be used for K value determination, as they enable broader ranges of K values to be covered with good accuracy. Improper procedure is, in our opinion, the main cause of discrepancies between K values reported in the literature for the same compound by different sources, or for different coating thicknesses. Financial support of Supelco, Varian and National Sciences and Engineering Research Council of Canada is acknowledged.References 1 Arthur, C. L., and Pawliszyn, J., Anal. Chem., 1990, 62, 2145. 2 Zhang, Z., and Pawliszyn, J., Anal. Chem., 1993, 65, 1843. 3 Arthur, C. L., Killam, L. M., Motlagh, S., Lim, M., Potter, D. W., and Pawliszyn, J., Environ. Sci. Technol., 1992, 26, 979. 4 Potter, D. W., and Pawliszyn, J., J. Chromatogr., 1992, 625, 247. 5 Wittkamp, B. L., and Tilotta, D. C., Anal. Chem., 1995, 67, 600. 6 Hawthorne, S. B., Miller, D. J., Pawliszyn, J., and Arthur, C.L., J. Chromatogr., 1992, 603, 185. 7 Arthur, C. L., Pratt, K., Motlagh, S., Pawliszyn, J., and Belardi, R. P., J. High Resolut. Chromatogr., 1992, 15, 741. 8 Langenfeld, J. J., Hawthorne, S. B., and Miller, D. J., Anal. Chem., 1996, 68, 144. 9 Nilsson, T., Ferrari, F., and Facchetti, S., in Proceedings of the 18th International Symposium on Capillary Chromatography, Riva del Garda (Italy), May 20–24, 1996, ed. Sandra, P., and Davos, G., IOPMS, Kortrijk, 1996, p. 618. 10 Potter, D., and Pawliszyn, J., Environ.Sci. Technol., 1994, 28, 298. 11 Chai, M., Arthur, C. L., Pawliszyn, J., Belardi, R. P., and Pratt, K. F., Analyst, 1993, 118, 1501. 12 Buchholz, K., and Pawliszyn, J., Environ. Sci. Technol., 1993, 27, 2844. 13 Buchholz, K., and Pawliszyn, J., Anal. Chem., 1994, 66, 160. 14 Schaefer, B., and Engewald, W., Fresenius’ J. Anal. Chem., 1995, 352, 535. 15 Boyd-Boland, A. A., and Pawliszyn, J., J. Chromatogr., 1995, 704, 163. 16 Eisert, R., Levsen, K., and Wuensch, G., J.Chromatogr., 1994, 683, 175. Table 2 Ranges of K values obtained when the amount of analyte extracted by the fiber, n, determined experimentally falls within ±5% (relative) of the true value, for two different fiber coating thicknesses and two sample volumes 100 mm— 2 ml sample 35 ml sample K Range of K K Range of K 100 105–95 100 105–95 1 000 1067–935 1 000 1051–949 10 000 12 537–8172 10 000 10 598–9413 100 000 H–36 190 100 000 115 748–86 928 1 000 000 H–55 072 1 000 000 4 700 000–492 593 10 000 000 H–58 104 10 000 000 H–923 611 100 000 000 H–58 426 100 000 000 H–1 012 177 7 mm— 2 ml sample 35 ml sample K Range of K K Range of K 100 105–95 100 105–95 1 000 1051–949 1 000 1050–950 10 000 10 574–9434 10 000 10 504–9496 100 000 112 903–88 785 100 000 105 422–94 622 1 000 000 3 500 000–558 824 1 000 000 1 093 750–913 462 10 000 000 H–1 187 500 10 000 000 17 500 000–6 785 714 100 000 000 H–1 338 028 100 000 000 H19 000 000 Analyst, October 1997, Vol. 122 108517 Popp, P., Kalbitz, K., and Oppermann, G., J. Chromatogr., 1994, 687, 133. 18 Lee, X., Kumazawa, T., Taguchi, T., Sato, K., and Suzuki, O., Hochudoku, 1995, 13, 122. 19 Eisert, R., and Levsen, K., Fresenius’ J. Anal. Chem., 1995, 351, 555. 20 Levsen, K., and Eisert, R., J. Am. Soc. Mass Spectrom., 1995, 6, 1119. 21 Graham, K. N., Sarna, L. P., Webster, G. R. B., Gaynor, J. D., and Ng, H. Y. F., J. Chromatogr., 1996, 725, 129. 22 Magdic, S., and Pawliszyn, J., J.Chromatogr., 1996, 723, 111. 23 Pan, L., Adams, M., and Pawliszyn, J., Anal. Chem., 1995, 67, 4396. 24 G�orecki, T., and Pawliszyn, J., Anal. Chem., 1996, 68, 3008. 25 Chen, J., and Pawliszyn, J., Anal. Chem., 1995, 67, 2530. 26 Boyd-Boland, A. A., and Pawliszyn, J., Anal. Chem., 1996, 68, 1521. 27 Hirata, Y., and Pawliszyn, J., J. Microcol. Sep., 1994, 6, 443. 28 Louch, D., Motlagh, S., and Pawliszyn, J., Anal. Chem., 1992, 64, 1187. 29 Arthur, C. L., Killam, L.M., Buchholz, K. D., and Pawliszyn, J., Anal. Chem., 1992, 64, 1960. 30 Motlagh, S., a Pawliszyn, J., Anal. Chim. Acta, 1993, 284, 265. 31 Langenfeld, J. J., Hawthorne, S. B., and Miller, D. J., Anal. Chem., 1996, 68, 144. 32 Martos, P., Saraullo, A., and Pawliszyn, J., Anal. Chem., 1997, 69, 402. 33 Namiesnik, J., J. Chromatogr., 1984, 300, 79. 34 Martos, P., and Pawliszyn, J., Anal. Chem., 1997, 69, 206. Paper 7/01303E Received February 25, 1997 Accepted June 4, 1997 1086 Analyst, October 1997, Vol. 122 Effect of Sample Volume on Quantitative Analysis by Solid-phase Microextraction Part 1. Theoretical Considerations Tadeusz G�orecki† and Janusz Pawliszyn* Department of Chemistry and Waterloo Centre for Groundwater Research, University of Waterloo, Waterloo, Ontario, Canada N2L 3G1 This paper discusses the effect of sample volume on the amount of analyte extracted from a sample by solid-phase microextraction (SPME) in two-phase (sample–fiber coating) and three-phase (sample–headspace–fiber coating) systems.Up-to-date knowledge is summarized, and new concepts are introduced. The effect of sample volume on quantification and precision of results can be neglected only in rare cases. The minimum sample volume which ensures that the amount extracted, n, is lower than 1% of the initial amount of the analyte present in the sample, as well as the volume for which exactly half of the initial amount of the analyte is extracted, have been calculated for both two- and three-phase systems.It is critical that the volumes of samples and standards are the same during analysis by SPME. Extraction kinetics in headspace analysis is dependent on the headspace capacity. If it is sufficiently large, the analyte is extracted almost exclusively from the gaseous phase, and equilibration can be very fast. On the other hand, this causes a significant loss of sensitivity. The effect of sample volume on the determination of the value of the partition coefficient, K, is also discussed. If the change in concentration of the analyte in the sample at equilibrium is not taken into account, erroneous results are obtained.Even when a proper procedure is used, there are practical limitations to the accuracy of the K value determination. Large sample volumes should always be used for K value determination, as they enable broader ranges of K values to be covered with good accuracy.Keywords: Solid-phase microextraction; sample volume; partition coefficient Solid-phase microextraction (SPME) was introduced in 1990.1 Since then, the interest in this method continues to grow very rapidly. SPME utilizes a small, fused-silica fiber coated with a polymeric stationary phase for analyte extraction from the matrix. The fiber is mounted for protection in a syringe-like device. The stationary phase can be a very viscous liquid (e.g., polydimethylsiloxane, PDMS) or a porous solid.For liquid phases, analytes are absorbed until an equilibrium is reached in the system. The amount extracted under these conditions is dependent on the partition coefficient between the sample and the coating. Sampling in SPME can be carried out directly from gaseous or liquid samples, or from the headspace over liquid or solid samples.2 SPME has been used for many applications, including the determination of substituted benzene compounds, 3–5 caffeine in beverages,6 volatile organic compounds in water,7–9 polyaromatic hydrocarbons and polychlorinated biphenyls,10 chlorinated hydrocarbons,11 phenols,12–14 pesticides, 15–22 and fatty acids,23 as well as lead and tetraethyllead.24 Although, initially, SPME was used in conjunction with GC only, its applicability has been broadened to LC25,26 and supercritical fluid chromatography (SFC).27 Theoretical aspects of SPME analysis have been studied in detail.Louch et al.28 developed the theory for two-phase systems (sample and fiber coating), and Zhang and Pawliszyn2 for three-phase systems (sample–headspace–coating).Practical aspects have been discussed in numerous papers, including those of Arthur et al.29 and Motlagh and Pawliszyn.30 Although the concepts behind SPME are relatively simple, analysis of the contents of some papers, as well as conversations with numerous users, indicate that certain aspects of SPME remain unclear or are misunderstood by many people, which can easily lead to confusion and frustration with the technique.A notorious example is the often large discrepancies between the partition coefficient (distribution constant) values reported for the same coatings and compounds by different groups, or even different researchers in the same group. These discrepancies have led Langenfeld et al.31 to propose the term ‘effective distribution constant’ to describe the partition coefficients observed for given sample–coating systems.The aim of this series of papers is to discuss several aspects of SPME analysis that might be directly responsible for the problems observed. In particular, the effect of sample volume on the amount of analyte extracted in two- and three-phase systems, as well as on determination of partition coefficients, K, will be discussed. Part 1 briefly summarizes the current knowledge and introduces several new concepts. Part 2 will present experimental verification of the ideas presented.Sample Volume versus Amount Extracted in Two-phase Systems In two-phase systems (gaseous sample–coating or liquid sample–coating) at equilibrium, the initial amount of the analyte present in the sample is distributed between the sample and the SPME fiber coating. The mass balance in such systems can be described in the following way:28 C0Vs = CHs Vs + CHf Vf (1) where C0 is the initial concentration of the analyte in the sample, Vs the sample volume, CHs the concentration of the analyte in the sample at equilibrium, CHf the concentration of the analyte in the fiber coating at equilibrium and Vf the volume of the coating.It should be emphasized that for liquid samples the above mass balance applies only to systems with no headspace, or for analytes whose Henry’s constants are so low that their amount in the sample headspace can be neglected. Partitioning between the sample and fiber coating is governed by the partition coefficient, K, also called the distribution constant: K C C = ¥ ¥ f s (2) † On leave from the Faculty of Chemistry, Technical University of Gda�nsk, Poland.Analyst, October 1997, Vol. 122 (1079–1086) 1079Combination of eqns. (1) and (2) and a few simple rearrangements yield the final expression describing the amount extracted by the SPME fiber at equilibrium, n:28 n KC V V V KV = + 0 s f s f (3) It is a common notion that the term KVf in the denominator of eqn. (3) is usually so small that it can be neglected, in which case the amount of analyte extracted by the fiber coating at equilibrium would be independent of sample volume and simply described by n = KC0Vf.While this may be true for analytes characterized by very low coating–sample partition coefficients or for samples of very large volumes, in most cases the above assumption is incorrect, which can lead to significant errors, as will be discussed in the following sections. The concentration of the analyte in the sample at equilibrium can be determined from a simple mass balance: C C V n V s s s ¥ = - 0 (4) Dividing both sides of eqn.(4) by C0 yields: C C C V n C V n C V KV V KV V V KV s s s s f s f s s f ¥ = - = - = - + = + 0 0 0 0 1 1 (5) Also, by definition C C n n s ¥ = 0 0 (6) where n is the amount of analyte extracted from a sample of volume Vs and n0 is the amount that would be extracted by the fiber from a sample of infinite volume (in which case no significant decrease in the concentration of the analyte in the sample would be observed at equilibrium).Fig. 1 presents the dependence of n/n0 on Vs for (a) a 100 mm fiber (Vf = 0.65 ml) and (b) a 30 mm fiber (Vf = 0.14 ml), for sample volumes of up to 1 l and K values ranging from 1000 to 1 000 000. This situation would be characteristic of air sampling from typical glass bulbs, and hydrocarbons from the gasoline range. According to Martos et al.,32 K Å 10orresponds to C7, while K Å 1 000 000 corresponds to C15.It is obvious from Fig. 1 that except for compounds with the lowest K values, sample volume has a very significant effect on the amount extracted by the fiber. Practical consequences of this are very important. Calibration of the fiber is usually performed by exposing it to a standard gaseous mixture generated in a glass bulb of known volume by the static method.33 The results of such calibrations are valid only for gaseous samples of exactly the same volume.If the fiber is exposed to ambient air (an unlimited supply of sample), the amount extracted by the fiber can be significantly larger compared with limited volume samples. On the other hand, if the containers used for grab sampling of air are smaller than the bulb used for calibration, the amount extracted will be smaller than expected from the calibration graph. In both cases, erroneous quantitative results will be obtained. It is obvious, therefore, that the same volume should be used for both the sample and the standard.Alternatively, a different calibration procedure can be used. Dynamic generation of standard gas mixtures provides unlimited volumes of gas standards, and thus can be used to calibrate the fiber for direct ambient air sampling. Alternatively, calibration based on retention parameters as described by Martos and Pawliszyn34 can be used for PDMS fibres. In this procedure no standards are necessary.Direct analysis of aqueous samples is usually performed by exposing the fiber to a small volume of sample contained in a vial. Fig. 2 presents similar dependences determined for sample volumes of up to 40 ml, and three coating thicknesses, viz., 100, 30 and 7 mm (Vf = 0.028 ml). It should be emphasized again that no headspace can be present in the vial if the two-phase system model is to be applied. The thin coating (7 mm) fibers are often used for direct sampling of semi-volatile compounds, since thicker coatings cause equilibration times to be excessively long. It is clear from Fig. 2 that the sample volume is an Fig. 1 Dependence of n/n0 on Vs for (a) 100 mm fiber (Vf = 0.65 ml), and (b) 30 mm fiber (Vf = 0.14 ml), for direct sampling from large volume samples (two-phase system). Fig. 2 Dependence of n/n0 on Vs for (a) 100 mm fiber (Vf = 0.65 ml), (b) 30 mm fiber (Vf = 0.14 ml), and (c) 7 mm fiber (Vf = 0.028 ml), for direct sampling from small volume samples (two-phase system). 1080 Analyst, October 1997, Vol. 122important parameter even for compounds with relatively small K values, especially when very small sample volumes (1 ml) are used. Moreover, headspace sampling is normaly used for volatile compounds with small K values and direct sampling is used for semi-volatile compounds with large K values. Hence, sample volume cannot usually be neglected even when the 7 mm coating is used. The course of the curves indicates that in some cases (steep part of the curve, e.g., K = 1000 for a 100 mm coating, Vs between 1 and 5 ml) small differences in sample volume can result in relatively large differences in the amount extracted, which adversely affects the precision of the determination.Care should be taken, therefore, to ensure that the sample volume during direct sampling is always the same. In view of the above discussion, it is interesting to establish what the sample volume should be in order for the amount extracted by the fiber to be insignificant compared with the amount remaining in the sample after extraction. Eqn.(3) can be used for this purpose. Let us assume that the amount extracted, n = CfVf, fulfils the above condition when it is smaller than, or equal to, 1% of the initial analyte amount in the sample, i.e., CHf Vf @ 0.01 C0Vs (7) The equation can be rearranged to yield: C C V V KV V f f s f s ¥ @ 0 0 01 0 01 @ @ . . (8) since, according to the initial assumption, C0 � CHs .Under the above conditions, eqn. (3) can be simplifid to n = KC0Vf (9) Simple rearrangement of eqn. (8) yields the final condition: VS ! 100KVf (10) For a 100 mm fiber, whose volume is approximately 0.65 ml, eqn. (10) can be written as: VS ! 0.065 K [ml] (11) It follows from eqn. (11) that even for analytes whose partition coefficient values are as low as 100, the minimum sample volume when using a 100 mm thick coating should be 6.5 ml if the effect of sample volume on the amount extracted is to be neglected.Similar reasoning can be applied to calculate the sample volume from which exactly 50% of the analyte is extracted, i.e., n = CfVf = 0.5 C0Vs. In this case 0 5 0 0 . C V KC V V V KV s s f s f = + (12) After a few rearrangements, the final condition is obtained: Vs 50% = KVf (13) or, for a 100 mm fiber, Vs = K 3 0.65 3 1023 [ml] (14) It follows from eqn. (14) that when 1 ml samples are analyzed with 100 mm fibers, 50% of the initial amount of the analyte will be extracted when the K value is approximately 1540.Typical partition coefficients of semi-volatile compounds are usually higher; it is obvious, therefore, that sample volume has a very significant effect on the amount extracted when small sample volumes are used. Sample Volume versus Amount Extracted in Three-phase Systems In most cases liquid samples are placed in vials with some headspace remaining inside. At equilibrium, by definition the chemical potentials of the analyte in all three phases (liquid sample–headspace–fiber coating) must be the same.2 A very important consequence of this fact is that the amount of the analyte extracted by the fiber at equilibrium in a three-phase system is the same independently of where the fiber is located, be it the headspace or the liquid.Consequently, in systems where a headspace is present, a different dependence should be used to calculate the amount of analyte extracted by the fiber regardless of where the fiber is located, viz:2 n KC V V KV K V V = + + 0 s f f hs hs s (15) where Khs is the headspace–liquid partition coefficient Khs = CH hs/CHs (16) (CH hs is the concentration of the analyte in the headspace at equilibrium), and Vhs is the headspace volume.Compared with a two-phase system described by eqn. (3), the difference is the additional term KhsVhs in the denominator of eqn. (4). For volatile compounds, Khs is usually close to 1, which means that headspace volume can be neglected only when it is close to zero (a two-phase system). Semi-volatile compounds have much lower values of Khs; therefore, the KhsVhs term may be negligibly small; however, such an assumption should always be verified.For obvious reasons the dependence of the amount of analyte extracted by the fiber on sample volume is much more complex in three-phase systems. However, it can be still dealt with relatively easily if the total volume of the system (sample plus headspace) remains constant.In practice, this will be the usual case, as the vial volume remains constant, and the headspace volume is the difference between the total volume and the sample volume. In such systems the concentration of an analyte in the sample at equilibrium is determined by the following mass balance: C C V C V n V s s hs hs s ¥ ¥ = - - 0 (17) Dividing both sides of eqn. (17) by C0 yields: C C C V C V n C V s hs hs s s ¥ ¥ = - - 0 0 0 1 (18) From the definition of Khs [eqn.(16)], CH hs can be replaced by KhsCHs , while n is described by eqn. (15), which yields C C K C V C V KV KV K V V s ks s hs s f f hs hs s ¥ ¥ = - - + + 0 0 1 (19) Simple rearrangements yield the final dependence: C C V KV K V V s s f hs hs s ¥ = + + 0 (20) Similarly to a two-phase system, Cs H/C0 = n/n0 when the headspace volume is constant. Fig. 3 illustrates the effect of headspace volume on the amount of analyte extracted in a system of constant volume (4, 15 and 40 ml—typical vial sizes) by a 100 mm fiber, typically used for the analysis of volatile compounds in sample headspace.To allow for better comparison of the results, the xaxis has been defined as the ratio of headspace volume to sample volume. The following values of partition coefficients have been used to calculate the course of the curves: K = 400, 2500 and 10, and Khs = 0.15, 0.7 and 1.24, respectively. These values correspond approximately to those for chloroform, 1,1,1-trichloroethane and carbon tetrachloride.It is interesting that n/n0 is always the largest for the dependence corresponding Analyst, October 1997, Vol. 122 1081to chloroform, which has the lowest Khs value, even though its K value is also the lowest. This is because, with such a combination of Khs and K, only a fraction of the analyte is present in the sample headspace; therefore, the equilibrium concentration of the analyte in the sample remains relatively high.The course of the two other curves is also interesting. For small headspace volumes, n/n0 is higher for the curve corresponding to carbon tetrachloride than for that corresponding to 1,1,1-trichloroethane, which is consistent with the values of their partition coefficients, K. However, as carbon tetrachloride has the largest Khs, n/n0 drops faster for this compound than for 1,1,1-trichloroethane, as a result of which the two lines cross each other.This illustrates the significance of headspace volume on the analytical results. It is also interesting that the largest relative increase in n/n0 ( Å 44%) when moving from small to large vials is observed for 1,1,1-trichloroethane, with the highest K value. For compounds with lower K values the relative increase in the amount extracted is much lower ( Å 16 and Å 12% for chloroform and carbon tetrachloride, respectively). It is, therefore, the combination of K and Khs for a given compound that determines the magnitude of the effect of the sample volume on the amount extracted in three-phase systems with headspace.Headspace volume can be the critical factor determining the precision of the results in three-phase systems. It is relatively easy to measure sample volume accurately. However, vials are not manufactured to have exactly the same volume. Wall thicknesses and bottom shapes may differ from vial to vial.Also, the shape of the septum in a closed vial can vary from concave to convex. All these factors will affect the total volume of the system, and, therefore, the headspace volume, which is the difference between the total volume and sample volume. As illustrated in Fig. 3, for compounds with large Khs, changing the headspace volume can significantly affect the amount extracted, especially in the range of headspace volumes usually applied (Vhs/Vs @ 1). Such differences (especially related to septum shape) are usually more pronounced in small vials; therefore, worse precision can usually be expected when using such vials for headspace sampling.Similarly to two-phase systems, it is possible to calculate the minimum sample volume necessary for the amount extracted by the fiber to be insignificant compared with the amount remaining in the sample after extraction. It is again assumed that the condition is fulfilled when less than 1% of the initial amount present in the sample is extracted by the fiber, i.e., n = CHf Vf @ 0.01 C0Vs.However, in this case it cannot be assumed that CHs � Co, because even if the amount extracted by the fiber constitutes only 1% of the initial amount of the analyte, a significant fraction of the analyte might be present in the headspace. Eqn. (15) will, therefore, be used to calculate the criterion: 001 0 0 . C V KC V V KV K V V s s f f hs hs s ! + + (21) The headspace volume, Vhs, can be expressed as a fraction of the sample volume, Vhs = aVs. Simple rearrangements yield the final criterion: V KV aK s f hs ! 99 1+ (22) Compared with the criterion for a two-phase system [eqn.(10)], there is an additional term in the denominator of eqn. (22), viz., aKhs. For non-zero headspace volumes this term is always greater than 0, which means that the sample volume fulfilling the criterion is smaller in three-phase systems than in two-phase systems. This is because, ultimately, the amount of analyte extracted by the fiber is determined by the concentration of the analyte in the liquid sample at equilibrium.This concentration will be lower in three-phase systems than in two-phase systems, since a significant fraction of the analyte may be present in the headspace of the sample. As a consequence, the sensitivity of headspace SPME can only be lower than, or equal to, that of direct SPME if the system is allowed to reach equilibrium.The higher sensitivity for headspace SPME reported sometimes (especially for compounds with relatively large K and Khs values) is clearly due to non-equilibrium conditions when sampling directly from liquid samples. This is not unusual in the light of the fact that equilibration times for direct sampling from water can be as long as a few hours (especially with inefficient agitation), and the increases in the amount of analyte extracted might not be noticeable if the experiment is not carried out for sufficiently long times.In addition, during prolonged sampling, analyte losses via adsorption onto the sample vial walls, absorption by the exposed parts of the silicone rubber septum (the Teflon lining of the septum has to be pierced to introduce the fiber; since the needle of the SPME device is blunt, the lining usually breaks open exposing a small area of silicone substrate), microbial decomposition, etc., can more than outweigh the expected increase in the amount of analyte extracted, as a result of which a smaller extracted amount can actually be observed after a longer time.This and other factors, including elimination of matrix effects related to the presence of high-boiling compounds in the sample and significantly faster equilibration times, usually make headspace SPME the preferred method for the analysis of mostly volatile compounds. It is also easy to determine the sample volume from which 50% of the analyte will be extracted by the SPME fiber placed in the sample headspace.The procedure is similar to that Fig. 3 Effect of headspace volume on the amount of analyte extracted in a system of a constant volume of (a) 4, (b) 15 and (c) 40 ml by a 100 mm fiber for chloroform, 1,1,1-trichloroethane and carbon tetrachloride (see text for the values of partition coefficients used for calculations). 1082 Analyst, October 1997, Vol. 122presented for two-phase systems. The final criterion in this case is: V KV aK s f hs 50 1 % = + (23) The difference between eqns.(23) and (13) is again the additional term in the denominator, aKhs. The higher the headspace–sample partition coefficient, Khs, or the headspaceto- sample volume ratio, a, the smaller the sample volume from which 50% of the initial amount of the analyte is extracted by the SPME fiber. Effect of Headspace Capacity on Extraction Kinetics Headspace volume can have a significant effect on equilibration times (extraction kinetics).If the headspace capacity is low (small Khs), and K is large, the equilibration process is very slow. A rapid initial increase in the amount of analyte extracted is usually observed, followed by a much slower increase that lasts for a long time. The first stage corresponds to analyte extraction from the gaseous phase only. As soon as the headspace concentration of the analyte falls below the equilibrium level with respect to the aqueous phase, analyte molecules start to move from the liquid sample to the headspace.At any given moment there can only be so many molecules in the headspace, depending on the Khs value, which causes the equilibration process to be very slow. The headspace acts in this case as a bottleneck for analyte transport to the fiber. On the other hand, if the amount of the analyte extracted by the fiber at equilibrium is negligible compared with the amount present in the headspace equilibrated with the sample, only a very small amount of the analyte has actually to be transported from the liquid sample through the headspace to the fiber coating, i.e., the analyte is extracted almost exclusively from the gaseous phase, and the process is much faster than for the case described above.Assuming this situation occurs when 95% of the analyte extracted by the fiber at equilibrium comes exclusively from the headspace, the criterion that must be fulfilled cabe described by eqn.(24). The assumption is reasonable, as a 5% difference usually falls within the limits of experimental error for trace SPME–GC analysis: R KV K V KV K aV = = f hs hs f hs s @ 0.05 (24) The above criterion means that the capacity of the headspace (KhsVhs) needs to be at least 20 times larger than the capacity of the fiber (KVf) to achieve rapid extraction. For a given sample volume, Vs, this can be achieved by using a sufficiently large headspace volume (corresponding to a large headspace-tosample volume ratio, a), or by increasing Khs.The latter can be accomplished by increasing the temperature or by salting the analyte out of the liquid phase. When the criterion described by eqn. (24) is fulfilled, equilibration can take as little as a few minutes, and is almost independent of the agitation conditions (provided that the analyte is equilibrated between the liquid phase and its headspace before the extraction begins). It should be emphasized, however, that great care must be exercised when moving the method from one vial size to another, as illustrated in Fig. 4. When the K value is large and/or Khs is small [Fig. 4(a)] a significant portion of the analyte extracted by the fiber has to be transported from the liquid sample through the headspace independently of the headspace volume, as evidenced by the values of R larger than 0.05 for all three values of a illustrated except for a = 2 and sample volumes larger than 28 ml (hence headspace volumes larger than 56 ml).The equilibration times will be relatively long for all vial sizes up to 40 ml. On the other hand, when the K value is smaller and/or Khs is larger, the R value can be lower than 0.05 for sample and headspace volumes often found in practice. Whether the criterion is fulfilled or not depends on the actual headspace volume [Fig. 4(b)]. For the same headspace-to-sample volume ratio, a, the criterion is fulfilled for large headspace volumes, but not for small ones.For example, when using a headspace-tosample volume ratio a = 0.5, the criterion is fulfilled for headspace volumes greater than 7 ml ( = sample volumes greater than 14 ml); therefore, under such conditions equilibration is very fast. However, when a 2 ml sample is used and the headspace volume is 1 ml, R is equal to 0.345, which indicates that a significant portion of the analyte has to be transported to the fiber from the liquid sample.As a consequence, the equilibration time will be longer even though the overall sample volume is smaller. If this is not accounted for and extraction times determined for large vials are used with small vials, the accuracy and precision of the determination suffers. It should be remembered, therefore, that the equilibration time has to be examined whenever the volume of the sample and/or headspace, or the extraction conditions (temperature, salt addition) are changed. It should also be emphasized that the increase in headspace capacity causes a significant loss of sensitivity.On the other hand, increasing the headspace capacity (which can be accomplished, for example, by increasing the extraction temperature) might dramatically shorten the equilibration times for some semi-volatile analytes while maintaining sufficient sensitivity. Both these effects need to be taken into account when developing a method involving headspace extraction. Effect of Sample Volume on the Determination of K in Two-phase Systems The fiber coating–sample partition coefficient [eqn.(2)] can be determined in several ways. The best way is to use eqn. (3). After a few rearrangements, the following dependence is obtained: K nV V C V n = - s f s ( ) 0 (25) Fig. 4 Dependence of R = KVf/KhsaVs on sample volume for three headspace-to-sample volume ratios, a = 0.5, 1 and 2, and a 100 mm PDMS fiber (Vf = 0.65 ml); (a) K = 1000, Khs = 0.25; (b) K = 250, Khs = 0.5.Analyst, October 1997, Vol. 122 1083If the amount extracted by the fiber, n, is negligible compared with the initial amount of the analyte in the sample, C0Vs, this equation is simplified to: K n C V = 0 f (26) which is a simple rearrangement of eqn. (9). Often, K is calculated directly from its definition [eqn. (2)]. In this case, if n is negligible compared with the initial amount of the analyte in the sample, C0Vs, then CHs can be substituted by C0, and the calculation is straightforward.However, it is our belief that often this substitution is made even when it is not legitimate. The consequences of this can be very significant, as will be illustrated in the following paragraphs. In both cases, the fiber volume necessary to calculate the concentration of the analyte in the fiber coating at equilibrium must be known with sufficiently good accuracy to obtain valid results. Let us introduce a new term, K apparent (Kap), this being the ratio of the concentration of the analyte in the fiber at equilibrium to the initial concentration C0: K C C ap f = ¥ 0 (27) Kap is what is calculated when depletion of the initial concentration of the analyte in the sample is not taken into account (i.e., when C0 is used instead of CHs ).The ratio of K to Kap is equal to: K K C C ap s = ¥ 0 (28) Therefore, K C C K ap s = � ¥ 0 (29) Since (5), s s s f C C V V KV ¥ = + 0 K KV V KV ap s s f = + (30) The true value of K can be calculated using the following equation derived from eqn.(29): K K V K V V = - - ap s ap f s (31) However, in many cases it will not be possible to determine K accurately by this method owing to the phenomena illustrated in Fig. 5. Fig. 5 presents the relationship between Kap and K for a wide range of partition coefficients, for two different coatings (100 and 7 mm) and three sample volumes (2, 10 and 30 ml). The dependences level off in all cases.It is obvious that the relationship reaches a certain limit, above which the same value of Kap is determined independently of the true value of K. The limit can be easily determined mathematically: lim lim K K K KV V KV V V ®¥ ®¥ = + æ è ç ö ø ÷ = ap s s f s f (32) It follows from eqn. (31) that Kap is determined only by the phase ratio Vs/Vf. The smaller the sample size (or the thicker the coating), the lower the value of Kap. It is possible that this phenomenon is behind certain very low K values reported in the literature for compounds that should have very large partition coefficients (e.g., polyaromatic hydrocarbons), especially when small sample volumes are used.Also, it is clear that when Kap is calculated instead of K, the values for thinner fiber coatings are much higher than those for thicker coatings (the difference in volume between a 100 mm fiber coating and a 7 mm coating is more than an order of magnitude). Such apparent discrepancies have also been reported in the literature.Even if proper methodology is used, owing to practical limitations it is not always possible to determine K accurately, as will be illustrated in the following paragraphs. Table 1 presents the amount of analyte extracted from a 2 ml sample and the corresponding per cent. increase in the amount extracted when going from a smaller K to a larger K for a 100 mm thick fiber coating and 1 mg l21 initial concentration, for different K values.For small K values the increase in the amount extracted when K increases is significant, although not linear as would be the case for infinite sample volumes. For large K values the increase in the amount extracted when K increases becomes insignificant and falls within the limits of experimental error. Table 2 illustrates the significance of this phenomenon. It presents the ranges of K values that are obtained when the amount of analyte extracted by the fiber, n, determined experimentally falls within ±5% (relative) of the true value, for two sample volumes (2 and 35 ml) and two fiber coating thicknesses (100 and 7 mm).A ±5% error can be assumed typical for trace analysis by SPME–GC. Fig. 5 Relationship between Kap and K for two different coatings (100 and 7 mm) and three sample volumes (2, 10 and 30 ml). Table 1 Amount of analyte extracted from a 2l sample and % increase in the amount extracted for a 100 mm thick fiber coating and 1 mg l21 initial concentration, for different K values K n/ng Increase (%) 100 0.0629540 — 1 000 0.4905660 717 10 000 1.5294118 212 100 000 1.9402985 28 1 000 000 1.9938650 2.8 10 000 000 1.9993848 0.28 100 000 000 1.9999385 0.3 1084 Analyst, October 1997, Vol. 122It is clear from Table 2 that practical limitations to K determination can be very severe, especially when thick fiber coatings and low sample volumes are used. For a 2 ml sample and a 100 mm thick fiber, the error of K determination for K values around 10 000 is around ±20% for a ±5% relative error of n determination.If the true K values are higher by an order of magnitude, the values determined experimentally can range from Å 36 000 to infinity, which is of no use. In practice, owing to experimental errors, for large K values the value of n found experimentally can be larger than the initial amount of analyte in the sample. In such cases, negative K values would be obtained, which, of course, have no physical meaning.It is necessary in such cases to assume that the amount extracted, n, is equal to the initial amount of analyte in the sample (quantitative extraction), in which case infinite K values are obtained. On the other hand, K values of Å 1 000 000 can be determined with reasonable accuracy when large sample volumes (35 ml) and thin fiber coatings (7 mm) are used. It is our belief that the above-described phenomena account for most of the discrepancies between K values reported in the literature for the same substances and coatings.Other possible sources of errors include limited solubility of some analytes in water, as well as losses of analytes due to volatilization (when headspace is present in the system), absorption by exposed parts of silicone rubber septa, adsorption on glass, or biodegradation. The first two phenomena are more probable for volatile analytes, while the remaining phenomena are more significant for semi-volatile compounds, since when K values are very high, equilibration times become very long.From a practical point of view, partition coefficients, K, should always be determined using large sample volumes (nothing can be gained by using small volumes). For semi-volatile compounds characterized by large K values, only thin coatings (7 mm for PDMS) are practical for the experimental determination of K. Above a certain level, the accurate determination of K becomes virtually impossible.Conclusions Sample volume plays an extremely important role in analysis by SPME. In order to avoid errors or poor precision, care should be taken to ensure that the volumes of samples and standard solutions used for calibration are the same. In headspace SPME, the combination of K and Khs determines the magnitude of the effect of sample volume on the amount extracted by the fiber, n. With the same headspace-to-sample volume ratio, the magnitude of the effect is greatest for compounds with low Khs.The minimum sample volume which ensures that n is lower than 1% of the initial amount of analyte present in the sample (which means that the change in concentration of the analyte in the sample at equilibrium is negligible) can easily be calculated for both two- and three-phase systems, and so is the volume for which exactly half of the initial analyte amount is extracted. Variable vial volumes might cause poorer precision in headspace sampling, as only the volume of the liquid sample can be accurately controlled.Extraction kinetics in headspace sampling is determined by the headspace capacity, KhsVhs. If it is sufficiently large compared with the fiber capacity, KVf, then the analyte is extracted almost exclusively from the headspace, and equilibration can be very fast. The headspace capacity is determined by the actual headspace volume rather than by the headspace-tosample volume ratio a; therefore, dramatically different equilibration times might be observed when scaling down the method from large to small vials, even though a remains constant.On the other hand, a large headspace capacity causes a significant loss of method sensitivity. Sample volume should always be taken into account when determining K values. If the change in concentration of the analyte in the sample at equilibrium is not taken into account, erroneous results are obtained. Even when a proper procedure is used, there are practical limitations to the accuracy of the K value determination.Large sample volumes should always be used for K value determination, as they enable broader ranges of K values to be covered with good accuracy. Improper procedure is, in our opinion, the main cause of discrepancies between K values reported in the literature for the same compound by different sources, or for different coating thicknesses. Financial support of Supelco, Varian and National Sciences and Engineering Research Council of Canada is acknowledged.References 1 Arthur, C. L., and Pawliszyn, J., Anal. Chem., 1990, 62, 2145. 2 Zhang, Z., and Pawliszyn, J., Anal. Chem., 1993, 65, 1843. 3 Arthur, C. L., Killam, L. M., Motlagh, S., Lim, M., Potter, D. W., and Pawliszyn, J., Environ. Sci. Technol., 1992, 26, 979. 4 Potter, D. W., and Pawliszyn, J., J. Chromatogr., 1992, 625, 247. 5 Wittkamp, B. L., and Tilotta, D. C., Anal. Chem., 1995, 67, 600. 6 Hawthorne, S. B., Miller, D. J., Pawliszyn, J., and Arthur, C. L., J. Chromatogr., 1992, 603, 185. 7 Arthur, C. L., Pratt, K., Motlagh, S., Pawliszyn, J., and Belardi, R. P., J. High Resolut. Chromatogr., 1992, 15, 741. 8 Langenfeld, J. J., Hawthorne, S. B., and Miller, D. J., Anal. Chem., 1996, 68, 144. 9 Nilsson, T., Ferrari, F., and Facchetti, S., in Proceedings of the 18th International Symposium on Capillary Chromatography, Riva del Garda (Italy), May 20–24, 1996, ed. Sandra, P., and Davos, G., IOPMS, Kortrijk, 1996, p. 618. 10 Potter, D., and Pawliszyn, J., Environ. Sci. Technol., 1994, 28, 298. 11 Chai, M., Arthur, C. L., Pawliszyn, J., Belardi, R. P., and Pratt, K. F., Analyst, 1993, 118, 1501. 12 Buchholz, K., and Pawliszyn, J., Environ. Sci. Technol., 1993, 27, 2844. 13 Buchholz, K., and Pawliszyn, J., Anal. Chem., 1994, 66, 160. 14 Schaefer, B., and Engewald, W., Fresenius’ J. Anal. Chem., 1995, 352, 535. 15 Boyd-Boland, A. A., and Pawliszyn, J., J. Chromatogr., 1995, 704, 163. 16 Eisert, R., Levsen, K., and Wuensch, G., J. Chromatogr., 1994, 683, 175. Table 2 Ranges of K values obtained when the amount of analyte extracted by the fiber, n, determined experimentally falls within ±5% (relative) of the true value, for two different fiber coating thicknesses and two sample volumes 100 mm— 2 ml sample 35 ml sample K Range of K K Range of K 100 105–95 100 105–95 1 000 1067–935 1 000 1051–949 10 000 12 537–8172 10 000 10 598–9413 100 000 H–36 190 100 000 115 748–86 928 1 000 000 H–55 072 1 000 000 4 700 000–492 593 10 000 000 H–58 104 10 000 000 H–923 611 100 000 000 H–58 426 100 000 000 H–1 012 177 7 mm— 2 ml sample 35 ml sample K Range of K K Range of K 100 105–95 100 105–95 1 000 1051–949 1 000 1050–950 10 000 10 574–9434 10 000 10 504–9496 100 000 112 903–88 785 100 000 105 422–94 622 1 000 000 3 500 000–558 824 1 000 000 1 093 750–913 462 10 000 000 H–1 187 500 10 000 000 17 500 000–6 785 714 100 000 000 H–1 338 028 100 000 000 H19 000 000 Analyst, October 1997, Vol. 122 108517 Popp, P., Kalbitz, K., and Oppermann, G., J. Chromatogr., 1994, 687, 133. 18 Lee, X., Kumazawa, T., Taguchi, T., Sato, K., and Suzuki, O., Hochudoku, 1995, 13, 122. 19 Eisert, R., and Levsen, K., Fresenius’ J. Anal. Chem., 1995, 351, 555. 20 Levsen, K., and Eisert, R., J. Am. Soc. Mass Spectrom., 1995, 6, 1119. 21 Graham, K. N., Sarna, L. P., Webster, G. R. B., Gaynor, J. D., and Ng, H. Y. F., J. Chromatogr., 1996, 725, 129. 22 Magdic, S., and Pawliszyn, J., J. Chromatogr., 1996, 723, 111. 23 Pan, L., Adams, M., and Pawliszyn, J., Anal. Chem., 1995, 67, 4396. 24 G�orecki, T., and Pliszyn, J., Anal. Chem., 1996, 68, 3008. 25 Chen, J., and Pawliszyn, J., Anal. Chem., 1995, 67, 2530. 26 Boyd-Boland, A. A., and Pawliszyn, J., Anal. Chem., 1996, 68, 1521. 27 Hirata, Y., and Pawliszyn, J., J. Microcol. Sep., 1994, 6, 443. 28 Louch, D., Motlagh, S., and Pawliszyn, J., Anal. Chem., 1992, 64, 1187. 29 Arthur, C. L., Killam, L. M., Buchholz, K. D., and Pawliszyn, J., Anal. Chem., 1992, 64, 1960. 30 Motlagh, S., and Pawliszyn, J., Anal. Chim. Acta, 1993, 284, 265. 31 Langenfeld, J. J., Hawthorne, S. B., and Miller, D. J., Anal. Chem., 1996, 68, 144. 32 Martos, P., Saraullo, A., and Pawliszyn, J., Anal. Chem., 1997, 69, 402. 33 Namiesnik, J., J. Chromatogr., 1984, 300, 79. 34 Martos, P., and Pawliszyn, J., Anal. Chem., 1997, 69, 206. Paper 7/01303E Received February 25, 1997 Accepted June 4, 1997 1086 Analyst, October 1997, Vol. 122

ISSN:0003-2654    

DOI:10.1039/a701303e

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Step-gradient Capillary Electrochromatography Melvin R. Euerby*a, Declan Gilligana, Christopher M. Johnsona and Keith D. Bartleb a Pharmaceutical and Analytical R & D, Astra Charnwood, Bakewell Road, Loughborough, Leicestershire, UK LE11 5RH b School of Chemistry, University of Leeds, Leeds, UK LS2 9JT The analytical benefits of using a step-gradient in capillary electrochromatography (CEC) are demonstrated. The application of step-gradient CEC to the analysis of six diuretics of widely differing lipophilicities was evaluated and shown to result in a marked reduction in the analysis time and an improvement in the peak shape for later-eluting lipophilic components.When the step-gradient approach was performed in an automated mode, the retention time RSD for repeated injections was below 1%. Keywords: Step-gradient; capillary electrochromatography; diuretic analysis; repeatability Interest in the field of capillary electrochromatography (CEC) is growing rapidly.This is evident from the ever-increasing number of research publications relating to this technique and the current commercialisation of equipment and capillaries suitable for CEC use. At present, commercially available capillary electrophoresis (CE) equipment which is capable of performing CEC is limited to simple isocratic CEC experiments; a considerable drawback is its inability to perform gradient elution CEC. In order to realise the full potential of CEC, it is necessary to develop the capacity of gradient elution for the separation of complex mixtures of widely differing lipophilicities as in HPLC.1,2 Several publications have reported encouraging results using gradient CEC with ‘home-built’ systems.3,4 This prompted us to investigate the feasibility of employing step-gradient CEC on commercially available CE equipment in order to widen the scope of current CEC applications. Experimental CEC was performed on a 1996 version Hewlett Packard HP3D (Cheadle Heath, UK) CE system capable of operating at a pressure of up to 12 bar. A 230 mm 3 50 mm id CEC capillary packed with 3 mm CEC Hypersil C18 material was kindly supplied by Hypersil (Runcorn, UK).Detection was at 210 nm with a 10 nm bandwidth and a 1 s rise time. CEC was performed using an applied voltage of 30 kV with 8 bar of pressure at each end of the capillary as described previously5,6 and a capillary temperature of 15 °C. The mobile phase consisted of different proportions of acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water. Electrokinetic injections (5 kV for 15 s) of the analytes [0.2 mg ml21 in acetonitrile–water (50 + 50)] were made.Results and Discussion The literature contains relatively few examples of true gradient CEC, i.e., electrodriven separations3 rather than voltage assisted pressure driven separations.4 These examples have utilised ‘home-built’ HPLC-based systems and to date have shown extremely encouraging results.Liao et al.7 have recently applied the concept of step-gradient CEC to achieve zone sharpening in the isocratic analysis of four polycyclic hydrocarbons. This involved introducing the sample electrokinetically onto a capillary previously conditioned with a lower organic content mobile phase. The mobile phase vials were exchanged manually on the ‘home-built’ CEC system for vials with a higher organic content, only then was the voltage applied and the separation commenced.However, for the technique to gain in popularity, the approach must be feasible on commercially available CE systems. The pressurised CE system used in this study possesses the capability to perform step-gradients in an automated sequence. The analysis is started with the initial mobile phase conditions; then, at a pre-defined time interval, the voltage is removed and the inlet and outlet buffer vials are exchanged for the final mobile phase conditions; the voltage is re-applied and the analysis continued. Finally, as part of the method, after the last peak has eluted, the voltage is terminated, the initial buffer vials are replaced and the voltage is re-applied to condition the capillary to the initial run conditions prior to the next analysis.This concept was investigated using a test mixture consisting of six diuretics of differing lipophilicities whose separation has previously been reported under isocratic CEC conditions.6 The mixture contained four polar diuretics [chlorothiazide (1), hydrochlorothiazide (2), chlorthalidone (3) and hydroflumethiazide (4)] which required a relatively low per cent.acetonitrile (MeCN) isocratic mobile phase composition to achieve acceptable resolution [see Fig. 1(b)] and two lipophilic diuretics [bendroflumethiazide (5) and bumetanide (6)] which required a significantly higher per cent. MeCN content for their elution within a reasonable analysis time [see Fig. 1(a)]. It can be seen from Fig. 1(a) and (b) that, in order to separate all six components within a reasonable analysis time, the initial mobile phase should correspond to MeCN–Na2HPO4 (0.050 mol dm23, pH 2.5)–water (40 + 20 + 40) and that the final composition should correspond to MeCN–Na2HPO4 (0.050 mol dm23, pH Fig. 1 Isocratic CEC separation of six commonly used diuretic compounds (1–6, see Table 1 for identity). For CEC conditions see under Experimental. (a) Isocratic mobile phase: acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (60 + 20 + 20). Linear velocity as measured by the retention of thiourea = 0.5 mm s21.For peak efficiencies see Table 1. (b) Isocratic mobile phase: acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (40 + 20 + 40). Linear velocity as measured by the retention of thiourea = 0.6 mm s21. For peak efficiencies see Table 1. Analyst, October 1997, Vol. 122 (1087–1088) 10872.5)–water (60 + 20 + 20).A mobile phase of pH 2.5 was essential in order to separate the components (both acids and neutral analytes) in an ion-suppressed mode. The initial conditions were held for 6.5 min, after which the voltage was removed and the buffer vials were exchanged for those containing the final mobile phase composition and the voltage was re-applied. After 10.75 min, the voltage was removed, the initial buffer vials were replaced, and the voltage was reapplied; these conditions were run for 7.75 min to re-establish the initial conditions prior to the next injection.As can be seen from Fig. 2, the elution time of the first four peaks was comparable to that observed in the isocratic run [Fig. 1(b)], indicating that they had not experienced the final mobile phase composition. The border between the two mobile phase compositions caused a baseline disturbance in the UV signal at approximately 13.5 min. The two later peaks eluted as two extremely sharp peaks with efficiencies in the region of 0.5 million plates per metre, suggesting that they had experienced a gradient/focusing effect (see Table 1).The use of the stepgradient resulted in a reduction of the analysis time from 35 to 16.5 min (25 min including the re-equilibration step) compared with isocratic CEC. Three consecutive runs of the partially optimised step-gradient method gave reproducible chromatography with RSDs for the retention times ranging between 0.48 and 1.00% (see Fig. 3). Analysis of variance showed that there was no statistical difference, at the 95% confidence level, between the peak efficiencies of the diuretics (2–4) using either isocratic or stepgradient CEC. The latter approach gave a statistically significant reduction in peak efficiency for chlorothiazide (1). However, this only corresponded to an 8% loss of the mean peak efficiency. Our encouraging findings are in sharp contrast to those of B�utehorn and Pyell,8 who reported a marked deterioration of chromatographic performance in their work on step-gradient micellar electrokinetic chromatography.They postulated that the marked peak broadening was attributable to the effects of the interface between the two mobile phases employed passing the solute zone, leading them to conclude that practical stepgradient micellar electrokinetic chromatography was noteasible. Our preliminary evaluation of step-gradient CEC using a commercially available CE system has shown extremely encouraging results in that we have proved that the concept is practicable and that compounds with widely differing lipophilicities can be analysed by CEC.The step-gradient CEC approach described in this paper is flexible, automated and repeatable. This work opens up the exciting prospect of the development of commercially available ‘new generation’ CE systems capable of performing continuous-gradient CEC. We thank H. Lomax and P. Ross (Hypersil Ltd.) for kindly supplying the CEC Hypersil C18 50 mm id capillary, and R.Sample (Hewlett-Packard Ltd.) for technical assistance with the HP3D CE system. References 1 Jandera, P., and Chur�acek, J., in Gradient Elution in Column Liquid Chromatography, Journal of Chromatography Library, vol. 31, Elsevier, Amsterdam, 1985. 2 Andreolini, F., and Trisciani, A., J. Chromatogr. Sci., 1990, 28, 54. 3 Yan, C., Dadoo, R., Zare, R. N., Rakestraw, D. J., and Anex, D. S., Anal. Chem., 1996, 68, 2726. 4 Behnke, B., and Bayer, E., J. Chromatogr. A, 1994, 680, 93. 5 Robson, M. M., Roulin, S., Shariff, S. M., Raynor, M. W., Bartle, K. D., Clifford, A. A., Myers, P., Euerby, M. R., and Johnson, C. M., Chromatographia, 1996, 43, 313. 6 Euerby, M. R., Gilligan, D., Johnson, C. M., Roulin, S., Bartle, K. D., and Myers, P., J. Microcol. Sep., in the press. 7 Liao, J. L., Chen, N., Ericson, C., and Hjerten, S., Anal. Chem., 1996, 68, 3468. 8 B�utehorn, U., and Pyell, U., Chromatographia, 1996, 43, 237.Paper 7/03175K Received May 8, 1997 Accepted June 6, 1997 Fig. 2 Step-gradient CEC separation of six commonly used diuretic compounds (1–6). CEC conditions as in Fig. 1, except that a step-gradient programme was used, viz., 0–6.50 min, acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (40 + 20 + 40); 6.50–17.25 min, 60 + 20 + 20; 17.25–25.00 min, 40 + 20 + 40. For peak efficiencies see Table 1. Fig. 3 Repeatability of step-gradient CEC separation of six commonly used diuretic compounds (1–6).CEC conditions as in Fig. 2. Table 1 Mean ‘apparent’ analyte peak efficiencies (n = 3) as a function of isocratic and step-gradient mobile phase composition [MeCN–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water]; for experimental details see legends of Figs. 1 and 2 ‘Apparent’ peak efficiency/plates per column Step-gradient: Isocratic: Isocratic: 40 + 20 + 40 ? Analyte 40 + 20 + 40 60 + 20 + 20 60 + 20 + 20 Chlorothiazide (1) 20 300 Co-elution 18 700 Hydrochlorothiazide (2) 21 100 Co-elution 19 900 Chlorthalidone (3) 20 700 Co-elution 19 800 Hydroflumethiazide (4) 21 500 Co-elution 21 340 Bendroflumethiazide (5) 28 900 31 100 209 000 Bumetanide (6) 23 400 42 100 111 500 1088 Analyst, October 1997, Vol. 122 Step-gradient Capillary Electrochromatography Melvin R. Euerby*a, Declan Gilligana, Christopher M. Johnsona and Keith D. Bartleb a Pharmaceutical and Analytical R & D, Astra Charnwood, Bakewell Road, Loughborough, Leicestershire, UK LE11 5RH b School of Chemistry, University of Leeds, Leeds, UK LS2 9JT The analytical benefits of using a step-gradient in capillary electrochromatography (CEC) are demonstrated.The application of step-gradient CEC to the analysis of six diuretics of widely differing lipophilicities was evaluated and shown to result in a marked reduction in the analysis time and an improvement in the peak shape for later-eluting lipophilic components. When the step-gradient approach was performed in an automated mode, the retention time RSD for repeated injections was below 1%.Keywords: Step-gradient; capillary electrochromatography; diuretic analysis; repeatability Interest in the field of capillary electrochromatography (CEC) is growing rapidly. This is evident from the ever-increasing number of research publications relating to this technique and the current commercialisation of equipment and capillaries suitable for CEC use.At present, commercially available capillary electrophoresis (CE) equipment which is capable of performing CEC is limited to simple isocratic CEC experiments; a considerable drawback is its inability to perform gradient elution CEC. In order to realise the full potential of CEC, it is necessary to develop the capacity of gradient elution for the separation of complex mixtures of widely differing lipophilicities as in HPLC.1,2 Several publications have reported encouraging results using gradient CEC with ‘home-built’ systems.3,4 This prompted us to investigate the feasibility of employing step-gradient CEC on commercially available CE equipment in order to widen the scope of current CEC applications.Experimental CEC was performed on a 1996 version Hewlett Packard HP3D (Cheadle Heath, UK) CE system capable of operating at a pressure of up to 12 bar. A 230 mm 3 50 mm id CEC capillary packed with 3 mm CEC Hypersil C18 material was kindly supplied by Hypersil (Runcorn, UK).Detection was at 210 nm with a 10 nm bandwidth and a 1 s rise time. CEC was performed using an applied voltage of 30 kV with 8 bar of pressure at each end of the capillary as described previously5,6 and a capillary temperature of 15 °C. The mobile phase consisted of different proportions of acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water. Electrokinetic injections (5 kV for 15 s) of the analytes [0.2 mg ml21 in acetonitrile–water (50 + 50)] were made.Results and Discussion The literature contains relatively few examples of true gradient CEC, i.e., electrodriven separations3 rather than voltage assisted pressure driven separations.4 These examples have utilised ‘home-built’ HPLC-based systems and to date have shown extremely encouraging results. Liao et al.7 have recently applied the concept of step-gradient CEC to achieve zone sharpening in the isocratic analysis of four polycyclic hydrocarbons. This involved introducing the sample electrokinetically onto a capillary previously conditioned with a lower organic content mobile phase.The mobile phase vials were exchanged manually on the ‘home-built’ CEC system for vials with a higher organic content, only then was the voltage applied and the separation commenced. However, for the technique to gain in popularity, the approach must be feasible on commercially available CE systems. The pressurised CE system used in this study possesses the capability to perform step-gradients in an automated sequence.The analysis is started with the initial mobile phase conditions; then, at a pre-defined time interval, the voltage is removed and the inlet and outlet buffer vials are exchanged for the final mobile phase conditions; the voltage is re-applied and the analysis continued. Finally, as part of the method, after the last peak has eluted, the voltage is terminated, the initial buffer vials are replaced and the voltage is re-applied to condition the capillary to the initial run conditions prior to the next analysis.This concept was investigated using a test mixture consisting of six diuretics of differing lipophilicities whose separation has previously been reported under isocratic CEC conditions.6 The mixture contained four polar diuretics [chlorothiazide (1), hydrochlorothiazide (2), chlorthalidone (3) and hydroflumethiazide (4)] which required a relatively low per cent. acetonitrile (MeCN) isocratic mobile phase composition to achieve acceptable resolution [see Fig. 1(b)] and two lipophilic diuretics [bendroflumethiazide (5) and bumetanide (6)] which required a significantly higher per cent. MeCN content for their elution within a reasonable analysis time [see Fig. 1(a)]. It can be seen from Fig. 1(a) and (b) that, in order to separate all six components within a reasonable analysis time, the initial mobile phase should correspond to MeCN–Na2HPO4 (0.050 mol dm23, pH 2.5)–water (40 + 20 + 40) and that the final composition should correspond to MeCN–Na2HPO4 (0.050 mol dm23, pH Fig. 1 Isocratic CEC separation of six commonly used diuretic compounds (1–6, see Table 1 for identity). For conditions see under Experimental. (a) Isocratic mobile phase: acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (60 + 20 + 20). Linear velocity as measured by the retention of thiourea = 0.5 mm s21. For peak efficiencies see Table 1. (b) Isocratic mobile phase: acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (40 + 20 + 40).Linear velocity as measured by the retention of thiourea = 0.6 mm s21. For peak efficiencies see Table 1. Analyst, October 1997, Vol. 122 (1087–1088) 10872.5)–water (60 + 20 + 20). A mobile phase of pH 2.5 was essential in order to separate the components (both acids and neutral analytes) in an ion-suppressed mode. The initial conditions were held for 6.5 min, after which the voltage was removed and the buffer vials were exchanged for those containing the final mobile phase composition and the voltage was re-applied. After 10.75 min, the voltage was removed, the initial buffer vials were replaced, and the voltage was reapplied; these conditions were run for 7.75 min to re-establish the initial conditions prior to the next injection.As can be seen from Fig. 2, the elution time of the first four peaks was comparable to that observed in the isocratic run [Fig. 1(b)], indicating that they had not experienced the final mobile phase composition. The border between the two mobile phase compositions caused a baseline disturbance in the UV signal at approximately 13.5 min. The two later peaks eluted as two extremely sharp peaks with efficiencies in the region of 0.5 million plates per metre, suggesting that they had experienced a gradient/focusing effect (see Table 1). The use of the stepgradient resulted in a reduction of the analysis time from 35 to 16.5 min (25 min including the re-equilibration step) compared with isocratic CEC.Three consecutive runs of the partially optimised step-gradient method gave reproducible chromatography with RSDs for the retention times ranging between 0.48 and 1.00% (see Fig. 3). Analysis of variance showed that there was no statistical difference, at the 95% confidence level, between the peak efficiencies of the diuretics (2–4) using either isocratic or stepgradient CEC. The latter approach gave a statistically significant reduction in peak efficiency for chlorothiazide (1).However, this only corresponded to an 8% loss of the mean peak efficiency. Our encouraging findings are in sharp contrast to those of B�utehorn and Pyell,8 who reported a marked deterioration of chromatographic performance in their work on step-gradient micellar electrokinetic chromatography. They postulated that the marked peak broadening was attributable to the effects of the interface between the two mobile phases employed passing the solute zone, leading them to conclude that practical stepgradient micellar electrokinetic chromatography was not feasible.Our preliminary evaluation of step-gradient CEC using a commercially available CE system has shown extremely encouraging results in that we have proved that the concept is practicable and that compounds with widely differing lipophilicities can be analysed by CEC. The step-gradient CEC approach described in this paper is flexible, automated and repeatable.This work opens up the exciting prospect of the development of commercially available ‘new generation’ CE systems capable of performing continuous-gradient CEC. We thank H. Lomax and P. Ross (Hypersil Ltd.) for kindly supplying the CEC Hypersil C18 50 mm id capillary, and R. Sample (Hewlett-Packard Ltd.) for technical assistance with the HP3D CE system. References 1 Jandera, P., and Chur�acek, J., in Gradient Elution in Column Liquid Chromatography, Journal of Chromatography Library, vol. 31, Elsevier, Amsterdam, 1985. 2 Andreolini, F., and Trisciani, A., J. Chromatogr. Sci., 1990, 28, 54. 3 Yan, C., Dadoo, R., Zare, R. N., Rakestraw, D. J., and Anex, D. S., Anal. Chem., 1996, 68, 2726. 4 Behnke, B., and Bayer, E., J. Chromatogr. A, 1994, 680, 93. 5 Robson, M. M., Roulin, S., Shariff, S. M., Raynor, M. W., Bartle, K. D., Clifford, A. A., Myers, P., Euerby, M. R., and Johnson, C. M., Chromatographia, 1996, 43, 313. 6 Euerby, M. R., Gilligan, D., Johnson, C. M., Roulin, S., Bartle, K. D., and Myers, P., J. Microcol. Sep., in the press. 7 Liao, J. L., Chen, N., Ericson, C., and Hjerten, S., Anal. Chem., 1996, 68, 3468. 8 B�utehorn, U., and Pyell, U., Chromatographia, 1996, 43, 237. Paper 7/03175K Received May 8, 1997 Accepted June 6, 1997 Fig. 2 Step-gradient CEC separation of six commonly used diuretic compounds (1–6). CEC conditions as in Fig. 1, except that a step-gradient programme was used, viz., 0–6.50 min, acetonitrile–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water (40 + 20 + 40); 6.50–17.25 min, 60 + 20 + 20; 17.25–25.00 min, 40 + 20 + 40. For peak efficiencies see Table 1. Fig. 3 Repeatability of step-gradient CEC separation of six commonly used diuretic compounds (1–6). CEC conditions as in Fig. 2. Table 1 Mean ‘apparent’ analyte peak efficiencies (n = 3) as a function of isocratic and step-gradient mobile phase composition [MeCN–Na2HPO4 (0.050 mol dm23, pH 2.5) buffer–water]; for experimental details see legends of Figs. 1 and 2 ‘Apparent’ peak efficiency/plates per column Step-gradient: Isocratic: Isocratic: 40 + 20 + 40 ? Analyte 40 + 20 + 40 60 + 20 + 20 60 + 20 + 20 Chlorothiazide (1) 20 300 Co-elution 18 700 Hydrochlorothiazide (2) 21 100 Co-elution 19 900 Chlorthalidone (3) 20 700 Co-elution 19 800 Hydroflumethiazide (4) 21 500 Co-elution 21 340 Bendroflumethiazide (5) 28 900 31 100 209 000 Bumetanide (6) 23 400 42 100 111 500 1088 Analyst, October 1997, Vo

ISSN:0003-2654    

DOI:10.1039/a703175k

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

On-column Indirect Photothermal Interference Detection for Capillary Zone Electrophoresis Yonggang Hua, Jieke Chenga and Yanzhuo Deng*b a Department of Chemistry, Wuhan University, Hubei 430072, China b Center of Analysis and Testing, Wuhan University, Hubei 430072, China An indirect photothermal interference detection method, which is based on the diffraction of a probe laser beam by a capillary tube, for detecting metal cations separated by capillary zone electrophoresis (CZE) is described.In this capillary photothermal interference detector, a 2 mW He–Ne probe laser is employed to provide the probe beam and an 18 mW He–Ne pump laser is used to supply the pump beam. Factors that contribute to the noise and signal in indirect photothermal interference detection in CZE, such as the noise from the scattered light caused by both the probe beam and the pump laser beam, and effects of the concentration of ethanol and NaCl on the separation efficiency, were studied.The effect of scattered light can be reduced effectively with the configuration constraints investigated for photothermal capillary absorption measurements. With Methylene Blue added to carrier electrolyte as an absorber, three typical metal cations (KI, CuII and AlIII) were separated using this system. For CuII, a 2.1 3 1027 mol l21 concentration detection limit (S/N = 2) without preconcentration and a 1.02 3 10217 mol detection limit as absolute amount were measured, considering the optical sampling volume estimated to be 50 pl, and 1.99 3 106 theoretical plates were observed on the laboratory-made CE system.It was demonstrated that indirect photothermal interference detection is suitable for small capillaries and a large ionic strength range for CE analysis. Keywords: Indirect photothermal interference detection; capillary zone electrophoresis; metal cations; Methylene Blue As a powerful technique for separations, the number of applications of capillary zone electrophoresis (CZE) has increased greatly in recent years.In principle, CZE is very suitable for small ionic compounds. However, the application of CZE to inorganic species, especially metal cations, is rare, in contrast to the separation of biological macromolecules.1,2 The reason is probably that many of these classes of compounds lack chromophores at useful wavelengths or have such low molar absorptivities as to preclude adequate sensitivity with absorption detection.Cation determinations by CZE are also hampered by coulombic interactions of the cations with the surfaces of the fused-silica capillaries commonly employed in CZE. These interactions can result in band broadening, which reduces both detectability and resolution. Methods adopting color reaction techniques to improve the LODs of metal cations with absorbance detection or laserinduced fluorescence detection can be easily devised. For example, 8-hydroxyquinoline-5-sulfonic acid (HQS2) was used for the determination of metal ions by CZE with laser-induced fluorescence.3 However, the main disadvantage is that the chemical properties of these complexes are usually unstable during the process of separation and it is difficult to optimize the separation and detection conditions for these complexes.4 The second disadvantage is that multiple complexes may exist in solution and multiple peaks would appear, making it very difficult to distinguish the signal peaks.The third disadvantage is that the analytes are chemically altered and collection and further studies are almost impossible. The fourth disadvantage is that derivatization is time consuming and inefficient. Hence there is a need for an all-purpose detector for CZE. Indirect detection is the most common method for metal cations which lack chromophores in CZE. A comprehensive account of indirect detection methods has been given by Yeung.5 Several reasons for developing indirect detection schemes have been mentioned.First, indirect detection is universal, and can be used for compounds which lack chromophores or fluorophores. Second, it is possible to broaden the applicability of highly sensitive but selective detectors by implementing indirect detection. Third, quantification is easier with indirect detection since tedious chemical derivatization procedures can be avoided. Fourth, indirect detection is nondestructive since no chemical manipulation is necessary and collection and further studies are facilitated.The indirect signal (displacement peaks) is a minor perturbation on the background signal. The concentration limit of detection, Clim, is given by6 C C TR DR lim = � M (1) where CM is the concentration of the relevant component added to the mobile phase, TR is the displacement ratio and DR is the dynamic reserve and is equal to the signal-to-noise ratio (S/N). It indicates that, for a given system, the more stable the background signal (larger DR), the more effective the displacement process (larger TR) and the lower is CM, the lower are the detection limits that can be obtained.Because DR and TR possibly decrease as one decreases CM, on the other hand, in order to improve the concentration LOD, detection methods with a capability for ultratrace detection and a larger range of detection are necessary to maintain a larger and stable background signal so as to allow the use of CM values that are as low as possible.Conductivity detection is an important indirect method for metal cations in CZE,7 which also functions in a displacement mode. The signal arises from the difference in equivalent conductance or mobility of the charge carrier electrolyte ion and the analyte ion. Several problems are usually encountered with conductivity detection. (1) It is advantageous to use a low ionic strength, poorly ionized buffer system to produce a low conductivity background.If good separation is desired, the concentration of the background electrolyte must be high relative to the concentration of the sample. However, this condition (high electrolyte concentration) results in a decrease in sensitivity due to the elevated background conductivity. (2) Background noise from the high voltage prevents highsensitivity detection. (3) A large difference between the mobility of the carrier electrolyte ion and that of the analyte ion leads to excessive peak tailing/fronting in CE. (4) Cell design is difficult since there must be no significant voltage drop between the cell electrodes; small dimensions of the capillary, high ionic strengths and a high separation electric field are challenges for the use of conductivity detection in CZE.Analyst, October 1997, Vol. 122 (1089–1093) 1089Indirect photometric detection, in which the signal arises from the difference in equivalent absorptivities between the carrier/eluent and the analyte ion, can offer a significant advantage over conductimetric detection in that ionic mobility and optical absorption are unrelated parameters.It is possible to choose carrier ions of suitable mobility and with desired optical absorption characteristics. Hence it is not necessary to compromise between separation efficiency and detection sensitivity. Since DR for photometry will degrade as optical pathlength, CM, or the absorption strengths decrease because incoherent light sources are used with low intensity and the presence of stray light in UV detection, indirect UV detection, commonly adopted in commercial CE instruments, is limited to the detection of inorganic ions in capillaries of @50 mm id without preconcentration.8 In indirect laser fluorimetry efficient rejection of stray light is made possible by the high collimated beam, so indirect laser fluorimetry can be used for capillaries of < 25 mm id with ultratrace detection. The advantages of indirect laser fluorimetry become apparent as one tries to decrease CM to improve the concentration LOD or mass LOD.9 However, fluorescence gives rise to a noisier baseline than does absorption, the end result being that the LOD is not any better than those for indirect UV detection and conductivity detection.The common problem encountered with the indirect detection methods mentioned above is that a low ionic strength of the buffer is needed r sensitive detection.However, several disadvantages arise under this condition. (1) The surface of the column wall will begin to affect detection because retention (e.g., via ion exchange or absorption on the column wall) can significantly alter the background electrolyte concentration. This surface effect can further reduce TR as CM is lowered and lead to a higher LOD, and the advantage of improving the concentration LOD by decreasing CM would be offset.10 (2) Low ionic strengths can result in distortion of the analyte zones and reduce the resolution.11 (3) It is impossible to maintain a stable baseline following an injection.(4) Interaction between the ions in the injected sample and the surface of the fused-silica capillary may be responsible for this effect at low ionic strengths. This effect would lead to considerable column-tocolumn variability.12 (5) Low ionic strengths preclude the use of low or high pH conditions, as H+ and OH2 ions, respectively, will dominate the displacement. This results in a decrease in TR and must be balanced to obtain the best sensitivity possible for the system.13 Therefore, CZE is applicable to the separation of inorganic ions, but their detection with high detection sensitivity, a low detection volume and a high ionic strength matrix remains a challenge.Lasers, with their unique spatial coherence, are ideally suited as light sources for capillary detection. Applications of lasers in CZE include fluorescence,14 refractive index,15 Raman16 and photothermal absorbance detection.17 The applications of photothermal absorbance detection for CE reported in recent years have been used mainly in analyses for amino acids18 and nucleosides and nucleotides19 with derivatization.The sensitivity of photothermal absorption techniques is 2–3 orders of magnitude higher than those of conventional detection methods. Since the absorption wavelength of analytes is required to match the emission wavelength of the pump beam well and the sensitivity of photothermal detection is related to the molar absorptivity of the analytes, these detectors are therefore not suitable for detecting non-absorbing analytes.This is one important reason why crossed-beam photothermal (CBPT) techniques have not been employed in CE for determinations of metal ions. In this paper, an indirect photothermal interference detection for detecting metal cations without derivatization after CE separation is described.Methylene Blue was chosen as a background absorber. The addition of ethanol and sodium chloride to the background electrolyte solution (BGES) can reduce the absorption of Methylene Blue and metal cations on the capillary wall, enhance the detector signal and improve the separation efficiency. Experimental Apparatus The indirect photothermal interference detector is shown in Fig. 1. A 2 mW He–Ne laser (Wuhan Institute of Laser Technology, Wuhan, China) produces a probe beam with a wavelength of 632.8 nm.A 46 mm focal length biconvex lens is used to focus the probe beam into the capillary tube. After traversing the capillary tube, the beam propagates to a photodiode (Semiconductor Factory, Wuhan University, Wuhan, China) with a 0.5 mm slit width. An M1000 He–Ne laser (Shanghai Institute of Laser Technology, Shanghai, China) produces an 18 mW pump laser beam. A model 194A mechanical chopper (EG&G, Princeton Applied Research, Princeton, NJ, USA) modulates the pump beam in a square wave.This beam is focused with a 10 mm focal length biconvex lens into the capillary tube. A three-axis translation stage holds the pump beam lens to provide complete freedom in the location of the pump beam waist, a two-axis stage is used to move the capillary, which is fixed firm on a special installation with a pinhole along the beam path of the probe laser, in the plane formed by the two laser beams, and a two-axis stage moves the detector across the probe beam profile.The interaction of the pump laser beam with the samples results in a refractive index change within the sample. This refractive index change causes a movement of the interference fringes and a change in the probe beam intensity which is synchronized with the pump beam modulation function. The modulated component of the probe beam intensity is measured by using an ND240 lock-in amplifier (Nanjing University Instrument Factory, Nanjing, China).A reference signal for the lock-in amplifier is generated within the chopper head. During the process of the electrophoresis, the signals from the lock-in amplifier are recorded by our SE 790 recorder (Australia). The capillary electrophoresis system was constructed in the laboratory. A high-voltage dc power supply (0–30 kV) (Shanghai, China) is employed. Electrical connections to the capillary are enclosed within a plastic box equipped with a safety interlock that acts to prevent accidental contact with the high voltage.A 60 cm long polymide-coated fused-silica capillary (Yongnian Optical Fiber Factory, Hebei, China) of 50 mm id and 320 mm od is used for electrophoresis. The polymer coating of the detection region is scraped away carefully. The distance from the injector to the detector is 50 cm. During the experiment, the optical breadboard is held at ground potential. Fig. 1 Schematic diagram of the detection system.PL, probe laser; M1 and M2, mirrors; L1 and L2, lenses; Ph1 and Ph2, pinholes; Ch, chopper; C, capillary; HL, heat laser; D, photodiode; LI, lock-in amplifier; R, recorder. 1090 Analyst, October 1997, Vol. 122Reagents All reagents were of analytical-reagent grade and solutions were prepared by dissolution in doubly distilled, de-ionized water. Oxides of metals were obtained from Shanghai Chemical (Shanghai, China). Samples were diluted by using the carrier electrolyte to avoid stacking and introduced into the capillary by using hydrostatic injection from a height of 10 cm.BGES were diluted with the same carrier electrolyte. All solutions and the buffer were filtered with 0.22 mm cut-off cellulose acetate filters before use. This filtration greatly reduces noise spikes and prevents blockage of the small capillaries. All data were obtained at room temperature (25 ± 1 °C). Results and Discussion Choice of Background Absorber The characteristics of the background absorber are very important for detection and separation with the utilization of indirect absorption detection in CZE.A higher molar absorptivity (ev) at the detection wavelength and a low CM will give a lower detection limit, as described by Wang and Hartwick.20 Five color reagents were obtained in order to assess their suitability as indirect background absorbers before separation. Experiments showed that the absorption wavelengths of pyrocatechol and Eriochrome Black T could match the pump laser beam (632.8 nm).However, these color reagents were unstable during the separation process and faded easily, because phenolic hydroxyl radicals are easily oxidized in an air environment. Victoria Blue B and Coomassie Brilliant Blue G 250 are not easily dissolved in aqueous solution. The solubility could be improved by adding ethanol, acetone or dimethylformamide to the electrolyte solution, but it was found that some small solid grains appeared on evaporation of the solvent, and these blocked the capillaries.Therefore, the color reagents mentioned above are not suitable for detection or separation in CZE. The molar absorptivity of Methylene Blue at 633 nm is about 4.5 3 104 l mol21 cm21. Experiments showed that characteristics of Methylene Blue are suitable for indirect interference detection and CZE separation. Noise Analysis It was found that the noise in indirect photothermal interference detection includes at least three sources: the detector, the color reagent and Joule heating. A significant effort was made to minimize the vibration of optical components. For example, the optical components are held with massive fixtures to a wall-damped optical table.When a laser beam strikes a capillary, a scattered light beam with many light and dark fringes can be observed, as described by Folestad et al.21 and Kerker and Matijewc22 and this scattered light beam is an important noise source in detection.First, portions of this scattered light, coming from the scattered light beam produced by the probe laser and the pump laser, could retroreflect back to both the probe laser source and pump beam laser source. This kind of retroreflection could cause fluctuations of the laser source and increase the noise level. Second, it was found that the noise level increases when the portion of scattered light coming from the scattered light beam produced by the pump laser hits the photodiode in the far-field after the capillary.This part of the scattered light is modulated by a mechanical chopper, and both the pump laser beam and probe laser beam are at the same wavelength in this system. In the present work, the probe laser beam was focused into the capillary at a right-angle, which tilted up to the capillary tube axis (a Å 89°). After being focused by a biconvex lens, the modulated pump laser beam was tilted slightly down into the capillary tube axis (b Å 89°).Further, the pump laser beam and the probe beam crossed obliquely at the capillary at a rightangle (g Å 89°) and two pinholes were used to extinguish the scattered light from other optical components, as shown in Fig. 1. It was demonstrated that the noise caused by the scattered light could be minimized successfully in this configuration. The reason is that the beams of scattered light in the plane of the interaction of the laser beam and the capillary were tilted to the capillary tube axis and not easy to retroreflect to the laser source and photodiode. As noted by Higashijima et al.23 the use of Methylene Blue in an acidic buffer leads to peak tailing characteristic of absorption of the positively charged dye on the capillary surface and would lead to higher noise.As shown later in Fig. 4, this type of absorption on the capillary surface could be minimized by adding ethanol to the electrolyte solutions and using an electrolyte solution of higher ionic strength.This phenomenon can be attributed to the fact that ethanol and a higher ionic strength can limit the dissociation of silanol on the capillary surface and the absorption of Methylene Blue. Further, ethanol and a higher ionic strength also enhanced the Methylene Blue detection signal, because the dn/dT of solutions containing ethanol and a higher ionic strength electrolyte is larger than that of aqueous solutions.24 This was beneficial for improving the sensitivity of indirect photothermal interference detection. It was also found that an excessively low concentration of the color reagent would lead to higher noise, as the ratio of the background absorbance to the noise somehow depends on the concentration of the color reagent and decreases simultaneously. 25 Excessive Joule heating should be avoided in CE as it prevents efficient heat dissipation to attain high efficiencies and causes thermal instability in the detection region.In general, as described in the Introduction, a low ionic strength of the buffers is required in indirect detection. However, an important effect was found in our experiments, namely that, in a certain range, an increase in the separation voltage in CZE did not decrease but actually improved the sensitivity of indirect photothermal interference detection. The reason is that the temperature rise in the detection region inside the capillary would maintain a stable level under the conditions of a stable voltage,26 and this kind of temperature increase of the background electrolyte could lead to an increase in the dn/dT value, as demonstrated by Franko and Tran.27 The contribution to noise from the Joule heating was found to be small under the condition of a stable voltage.Indirect Detection Before detection for CZE separation, several alignment constraints were qualitatively investigated for photothermal capillary absorption measurements.As expected, the signal was highest when the pump laser beam waist was centered in the capillary. Since a tightly focused pump beam also minimizes the system volume, all measurements reported in this paper were obtained with the pump waist located in the capillary. For a given interference pattern, the fringes that were found to be more sensitive were those that appear near the optical axis of the probe beam but still retain high intensity and contrast.When the interference pattern and the slit width of detector were fixed, an optimum distance for detection from the sample cuvette to the photodiode was observed. Agreeing well with a lower noise level and higher sensitivity, in the experiment the photodiode was set at a position of 60 cm in the far-field after the capillary. Under these conditions, the capillary was located at a position of 19 mm before the probe beam waist to obtain higher sensitivity.When the capillary containing the solutions was drawn gradually closer to the probe beam waist, it was found that the signal decreased quickly. The reason may be that the contrast of the interference fringes also decreased quickly. Analyst, October 1997, Vol. 122 1091A finite time is required to reach thermal equilibrium in photothermal absorbance measurements. As a general rule of thumb, a frequency of 6.5 Hz was used in these experiments. A short time constant is of value since very fast phenomena may be studied without degradation due to instrumental artifacts, and the detector time constant was set at 1 s.An absorbance calibration curve of Methylene Blue was obtained in the static state for the 50 mm id capillaries. The calibration curve was linear (r = 0.995) over more than three orders of magnitude of concentration. Linearity extended from the detection limit, A = 6.1 31027, to the highest concentration sample studied, A = 0.023.Water–ethanol solution was chosen as the solvent because it produces greater sensitivity than water for photothermal absorbance measurements. As shown in Fig. 2, for Methylene Blue solution, the concentration LOD was about 3.1 3 1028 mol l21 (S/N = 2). With a highly collimated beam, the detection volume of this detector, the intersection volume of the pump and probe beam, is small, slightly less than 1 pl. We use a much more conservative definition of the detection volume, as described by Bornhop and Dovichi,28 as a cylinder whose radius and height are given by the capillary radius.For the 50 mm id tube, the detector volume was about 50 pl. Then, the detection limit as absolute amount was 1.5 3 10218 mol. Of course, a significantly smaller amount of analyte was present within the small intersection volume of the laser beam than the result estimated above. It has been demonstrated that, for absorption measurement in narrow-bore capillaries, crossed-beam photothermal (CBPT) absorption techniques can offer definite advantages: the sensitivities are not linear with respect to pathlength, and with a decrease in the capillary diameter the pump beam is more efficiently quenched at the walls.Therefore, according to the assumption described in ref. 28, this detector could also be used with a capillary tube as small as 5 mm id. The results in Fig. 3 show that the addition of ethanol to the BGES can reduce absorption on the capillary surface and lead to an improvement in the peak shape of the analyte.Ethanol also enhanced the Methylene Blue and analyte detection signal, because the dn/dT value of ethanolic solutions is larger than that of aqueous solutions, and this is beneficial for the improvement of sensitivity in indirect interference detection. Further, it could be observed that the elution time became longer with an increase of ethanol concentration. A probable cause is that the ethanol interacts strongly with the capillary wall, resulting in a higher apparent concentration of ethanol within the double layer as described in ref. 29. These interactions then result in higher apparent viscosities within the double layer and lead to a reduced electroosmotic flow. The effect of NaCl on CZE separation and detection was investigated. The results (Fig. 4) showed that efficiency of KI and the peak shape were improved at higher concentrations of NaCl without a decrease in detection sensitivity.These results and that of the Joule heating phenomenon (see above) indicated that indirect photothermal interference detection is suitable for CE separation systems with a high ionic strength buffer. As the sensitivity that CBPT provides was almost unrelated to the light path, this result implies that indirect photothermal interference detection may be used for detection in CZE with a larger ionic strength range and overcomes the problem of the decrease in sensitivity for small id capillaries and shot-noise limitation in photometry.The characteristics of photothermal absorbance detection and the results obtained above are, for example, beneficial for the analysis of samples with weak absorption, ultratrace detection and larger range of linearity, and permit a lower concentration of the background absorber (Methylene Blue) to be used to improve the LOD. As shown in Fig. 5, positive peaks are observed for KI and CuII and AlIII produces a negative peak.A positive peak indicates an increase in concentration of the background absorber present at the detector, whereas a negative peak indicates a decrease. The migration time of the system peak corresponds to the mobility of the Methylene Blue cation. An interesting phenomenon is observed in Fig. 5, namely that the migration order is KI > CuII > AlIII. In order to understand the elution order, it has to be taken into account that the electrophoretic mobility depends on the charge, shape and size of the analyte.This phenomenon could be explained by the fact that, under conditions of low pH and excess of chloride, Fig. 2 Electropherogram of Methylene Blue. Carrier electrolyte, 50 mmol l21 Na2HPO4–10% ethanol (pH 6.0); sample, 5 3 1027 mol l21 Methylene Blue; CE voltage, 16 kV; current, 14 mA; injection, 10 s. Fig. 3 Electropherograms of KI (1.0 3 1025 mol l21). Carrier electrolyte; (a) 109 mmol l21 acetic acid–5% ethanol; (b) 109 mmol l21 acetic acid 210% ethanol; and (c) 109 mmol l21 acetic acid–20% ethanol.CE voltage, 14 kV; injection, 15 s; Methylene Blue concentration, 5.0 3 1024 mol l21. Fig. 4 Electropherogram of KI (1.0 3 1025 mol l21). Carrier electrolyte, 109 mmol l21 acetic acid–20% ethanol–100 mmol l21 NaCl; CE voltage, 14 kV; current, 24 mA; other conditions as in Fig. 3. 1092 Analyst, October 1997, Vol. 122CuCl4 22 and AlCl42 complexes are formed in solution, but KI is incapable of forming chloride complexes.Therefore, the peak of KI appeared first. The elution order of CuCl4 22 and AlCl42 can be rationalized by the fact that AlCl42 seems to be a poorly hydrated anion compared with CuCl4 22. Hence this lower hydration will reduce the size of the moving ion, which may result in greater electrophoretic migration and, accordingly, retardation. The phenomenon is similar to the studies of Aguilar et al.30 and Baraj et al.,31 in which chloride and cyanide were used as ligands for the determination of metal cations.For CuII, the samples and carrier solution were prepared with almost the same conductivity, and consequently the stacking effect was not used; a 2.1 3 1027 mol l21 concentration detection limit (S/N = 2) without preconcentration and a 1.20 3 10217 mol detection limit as absolute amount, considering the optical sampling volume estimated to be 50 pl, were measured, and 1.99 3106 theoretical plates were observed with the laboratorymade CE system.The concentration LODs measured here were comparable to the best results obtained with other detection methods for ions,32–34 and the mass LOD of this detector could be decreased to that of an indirect fluorescence detector or conductivity detector (10217 mol) and at least two orders of magnitude lower than that of a commercial UV detector. The CE method offers several advantages over other indirect detection methods for ions. First the use of a larger range of ionic strengths permits the application dynamic range to be increased35,36 and extends the upper limit of sample ionic strength that still permits effective stacking and thus facilitates the determination of trace constituents in a high ionic strength matrix.Second, the lower mass LOD permits small id capillaries to be used to improve the resolution of samples and ultra trace components. Obviously, this characteristic of indirect photothermal interference detection is better than that of commercial detection.Even then the best mass LODs of conductimetric detection and indirect fluorescence detection are similar to those of this method; as one mode of absorption detection techniques, this method has all the advantages of photometric detection over conductimetric detection mentioned earlier, and maybe a less noisy baseline than that in indirect fluorescence detection, since the main disadvantage of indirect fluorescence detection is that fluorescence gives rise to a noisier baseline than does absorption and leads to a higher concentration LOD.Third, as the values of dn/dT for most non-aqueous media are larger than those of aqueous solutions, this detector will be of advantage for detection in non-aqueous medium capillary electrophoresis, an important field in the study of capillary electrophoresis. In future work we will investigate possible improvements to the detection sensitivity, including the design of an electronic noise canceller to reduce electronic noise, altering the injection mode and the use of highly stable lasers.The relatively low cost, simple construction, small volume, universal detection, larger range of ionic strengths and excellent detection limits of this method should prove attractive for a number of applications in capillary electrophoresis separation techniques. This work was supported by the National Nature Science Foundation of China. References 1 Stoker, F.S., Haymor, B. L., and McBeath, R., J. Chromatogr., 1989, 470, 241. 2 Burlity, N., and Jorgenson, J., J. Chromatogr., 1989, 480, 301. 3 Swaile, D. F., and Sepaniak, M. J., Anal. Chem., 1991, 63, 179. 4 Hu, Y., Deng, Y., and Cheng, J., Prog. Nat. Sci., 1996, 6, S-42. 5 Yeung, E. S., Acc. Chem. Res., 1989, 22, 125. 6 Yeung, E. S., and Kuhr, W. G., Anal. Chem., 1991, 63, 275A. 7 Kaniansky, D., Zelenska, V., and Baluchova, D., Electroporesis, 1996, 17, 1890. 8 Jandik, P., and Bonn, G., Capillary Electrophoresis of Small Ions, VCH, Weinheim, 1993. 9 Pfeffer, W. D., and Yeung, E. S., J. Chromatogr., 1990, 506, 401. 10 Wilson, S. A., and Yeung, E. S., Anal. Chim. Acta, 1984, 157, 53. 11 Mikkers, F. E. P., Everaerts, F. M., and Verheggen, Th. P. E. M., J. Chromatogr., 1979, 169, 11. 12 Kuhr, W. G., and Yeung, E. S., Anal. Chem., 1988, 60, 2642. 13 Takeuchi, T., and Yeung, E. S., J. Chromatogr., 1986, 370, 83. 14 Gussman, E., Kuo, R. N., and Zare, R. N., Science, 1985, 230, 813. 15 Pawliszyn, J., J. Liq. Chromatogr., 1987, 10, 3377. 16 Cheng, C. Y., and Morris, M. D., Appl. Spectrosc., 1988, 42, 515. 17 Yu, M., and Dovich, N. J., Mikrochimica III, 1988, 27. 18 Yu, M., and Dovich, N. J., Anal. Chem., 1989, 61, 37. 19 Krattiger, B., Bruno, A. E., Widmer, H. M., and Pandliker, R., Anal. Chem., 1995, 67, 124. 20 Wang, T. S., and Hartwick, R. A., J. Chromatogr., 1992, 607, 119. 21 Folestad, S., Johnson, L., Josefsson, B., and Galle, B., Anal.Chem., 1982, 54, 925. 22 Kerker, M., and Matijewc, E., J. Opt. Soc. Am., 1961, 51, 506. 23 Higashijima, T., Fuchigami, T., Imasaka, T., and Ishibashi, N., Anal. Chem., 1992, 64, 711. 24 Fujiwara, K., Lei, W., Uckiki, H., Shimokoshi, F., Fuwa, K., and Kobayashi, T., Anal. Chem., 1982, 54, 2026. 25 Takenchi, T., and Yeung, E. S., J. Chromatogr., 1986, 370, 83. 26 Bruno, A. E., Krattiger, B., Maystre, F., and Widmer, H. M., Anal. Chem., 1991, 63, 2689. 27 Franko, M., and Tran, C.D., J. Phys. Chem., 1991, 95, 6688. 28 Bornhop, D. J., and Dovichi, N. J., Anal. Chem., 1987, 59, 1632. 29 VanOrman, B. B., Liversidge, G. G., and McIntire, G. L., J. Microcolumn Sep., 1991, 2, 176. 30 Aguilar, M., Farran, A., and Martinnez, M., J. Chromatogr., 1993, 635, 127. 31 Baraj, B., Sastre, A., Merkoci, A., and Martinnez, M., J. Chromatogr., 1995, 718, 227. 32 Weston, A., Broun, P. R., Jandik, P., Heckenberg, A. L., and Jones, W. R., J. Chromatogr., 1992, 608, 395. 33 Kuhr, W. G., and Yeung, E. S., Anal. Chem., 1988, 60, 2642. 34 Kaniansky, D., Zelenska, V., and Baluchova, D., Electrophoresis, 1996, 17, 1890. 35 Nielen, M. W. F., J. Chromatogr., 1991, 542, 173. 36 Green, J. S., and Jorgenson, J. W., J. Chromatogr., 1987, 478, 63. Paper 7/00119C Received January 6, 1997 Accepted June 18, 1997 Fig. 5 Electropherogram of a mixture of three metal cations. Methylene Blue concentration, 5.0 3 1026 mol l21. Other conditions as in Fig. 4. Peaks: 1, KI (2.5 3 1026 mol l21); 2, system peak; 3, CuII (1.56 3 1026 mol l21); and 4, AlIII (3.85 3 1026 mol l21). Analyst, October 1997, Vol. 122 1093 On-column Indirect Photothermal Interference Detection for Capillary Zone Electrophoresis Yonggang Hua, Jieke Chenga and Yanzhuo Deng*b a Department of Chemistry, Wuhan University, Hubei 430072, China b Center of Analysis and Testing, Wuhan University, Hubei 430072, China An indirect photothermal interference detection method, which is based on the diffraction of a probe laser beam by a capillary tube, for detecting metal cations separated by capillary zone electrophoresis (CZE) is described.In this capillary photothermal interference detector, a 2 mW He–Ne probe laser is employed to provide the probe beam and an 18 mW He–Ne pump laser is used to supply the pump beam. Factors that contribute to the noise and signal in indirect photothermal interference detection in CZE, such as the noise from the scattered light caused by both the probe beam and the pump laser beam, and effects of the concentration of ethanol and NaCl on the separation efficiency, were studied.The effect of scattered light can be reduced effectively with the configuration constraints investigated for photothermal capillary absorption measurements. With Methylene Blue added to carrier electrolyte as an absorber, three typical metal cations (KI, CuII and AlIII) were separated using this system.For CuII, a 2.1 3 1027 mol l21 concentration detection limit (S/N = 2) without preconcentration and a 1.02 3 10217 mol detection limit as absolute amount were measured, considering the optical sampling volume estimated to be 50 pl, and 1.99 3 106 theoretical plates were observed on the laboratory-made CE system. It was demonstrated that indirect photothermal interference detection is suitable for small capillaries and a large ionic strength range for CE analysis. Keywords: Indirect photothermal interference detection; capillary zone electrophoresis; metal cations; Methylene Blue As a powerful technique for separations, the number of applications of capillary zone electrophoresis (CZE) has increased greatly in recent years.In principle, CZE is very suitable for small ionic compounds. However, the application of CZE to inorganic species, especially metal cations, is rare, in contrast to the separation of biological macromolecules.1,2 The reason is probably that many of these classes of compounds lack chromophores at useful wavelengths or have such low molar absorptivities as to preclude adequate sensitivity with absorption detection. Cation determinations by CZE are also hampered by coulombic interactions of the cations with the surfaces of the fused-silica capillaries commonly employed in CZE.These interactions can result in band broadening, which reduces both detectability and resolution. Methods adopting color reaction techniques to improve the LODs of metal cations with absorbance detection or laserinduced fluorescence detection can be easily devised.For example, 8-hydroxyquinoline-5-sulfonic acid (HQS2) was used for the determination of metal ions by CZE with laser-induced fluorescence.3 However, the main disadvantage is that the chemical properties of these complexes are usually unstable during the process of separation and it is difficult to optimize the separation and detection conditions for these complexes.4 The second disadvantage is that multiple complexes may exist in solution and multiple peaks would appear, making it very difficult to distinguish the signal peaks.The third disadvantage is that the analytes are chemically altered and collection and further studies are almost impossible. The fourth disadvantage is that derivatization is time consuming and inefficient. Hence there is a need for an all-purpose detector for CZE. Indirect detection is the most common method for metal cations which lack chromophores in CZE.A comprehensive account of indirect detection methods has been given by Yeung.5 Several reasons for developing indirect detection schemes have been mentioned. First, indirect detection is universal, and can be used for compounds which lack chromophores or fluorophores. Second, it is possible to broaden the applicability of highly sensitive but selective detectors by implementing indirect detection.Third, quantification is easier with indirect detection since tedious chemical derivatization procedures can be avoided. Fourth, indirect detection is nondestructive since no chemical manipulation is necessary and collection and further studies are facilitated. The indirect signal (displacement peaks) is a minor perturbation on the background signal. The concentration limit of detection, Clim, is given by6 C C TR DR lim = � M (1) where CM is the concentration of the relevant component added to the mobile phase, TR is the displacement ratio and DR is the dynamic reserve and is equal to the signal-to-noise ratio (S/N).It indicates that, for a given system, the more stable the background signal (larger DR), the more effective the displacement process (larger TR) and the lower is CM, the lower are the detection limits that can be obtained. Because DR and TR possibly decrease as one decreases CM, on the other hand, in order to improve the concentration LOD, detection methods with a capability for ultratrace detection and a larger range of detection are necessary to maintain a larger and stable background signal so as to allow the use of CM values that are as low as possible.Conductivity detection is an important indirect method for metal cations in CZE,7 which also functions in a displacement mode. The signal arises from the difference in equivalent conductance or mobility of the charge carrier electrolyte ion and the analyte ion.Several problems are usually encountered with conductivity detection. (1) It is advantageous to use a low ionic strength, poorly ionized buffer system to produce a low conductivity background. If good separation is desired, the concentration of the background electrolyte must be high relative to the concentration of the sample. However, this condition (high electrolyte concentration) results in a decrease in sensitivity due to the elevated background conductivity.(2) Background noise from the high voltage prevents highsensitivity detection. (3) A large difference between the mobility of the carrier electrolyte ion and that of the analyte ion leads to excessive peak tailing/fronting in CE. (4) Cell design is difficult since there must be no significant voltage drop between the cell electrodes; small dimensions of the capillary, high ionic strengths and a high separation electric field are challenges for the use of conductivity detection in CZE.Analyst, October 1997, Vol. 122 (1089–1093) 1089Indirect photometric detection, in which the signal arises from the difference in equivalent absorptivities between the carrier/eluent and the analyte ion, can offer a significant advantage over conductimetric detection in that ionic mobility and optical absorption are unrelated parameters. It is possible to choose carrier ions of suitable mobility and with desired optical absorption characteristics. Hence it is not necessary to compromise between separation efficiency and detection sensitivity.Since DR for photometry will degrade as optical pathlength, CM, or the absorption strengths decrease because incoherent light sources are used with low intensity and the presence of stray light in UV detection, indirect UV detection, commonly adopted in commercial CE instruments, is limited to the detection of inorganic ions in capillaries of @50 mm id without preconcentration.8 In indirect laser fluorimetry efficient rection of stray light is made possible by the high collimated beam, so indirect laser fluorimetry can be used for capillaries of < 25 mm id with ultratrace detection.The advantages of indirect laser fluorimetry become apparent as one tries to decrease CM to improve the concentration LOD or mass LOD.9 However, fluorescence gives rise to a noisier baseline than does absorption, the end result being that the LOD is not any better than those for indirect UV detection and conductivity detection.The common problem encountered with the indirect detection methods mentioned above is that a low ionic strength of the buffer is needed for sensitive detection. However, several disadvantages arise under this condition. (1) The surface of the column wall will begin to affect detection because retention (e.g., via ion exchange or absorption on the column wall) can significantly alter the background electrolyte concentration. This surface effect can further reduce TR as CM is lowered and lead to a higher LOD, and the advantage of improving the concentration LOD by decreasing CM would be offset.10 (2) Low ionic strengths can result in distortion of the analyte zones and reduce the resolution.11 (3) It is impossible to maintain a stable baseline following an injection.(4) Interaction between the ions in the injected sample and the surface of the fused-silica capillary may be responsible for this effect at low ionic strengths.This effect would lead to considerable column-tocolumn variability.12 (5) Low ionic strengths preclude the use of low or high pH conditions, as H+ and OH2 ions, respectively, will dominate the displacement. This results in a decrease in TR and must be balanced to obtain the best sensitivity possible for the system.13 Therefore, CZE is applicable to the separation of inorganic ions, but their detection with high detection sensitivity, a low detection volume and a high ionic strength matrix remains a challenge.Lasers, with their unique spatial coherence, are ideally suited as light sources for capillary detection. Applications of lasers in CZE include fluorescence,14 refractive index,15 Raman16 and photothermal absorbance detection.17 The applications of photothermal absorbance detection for CE reported in recent years have been used mainly in analyses for amino acids18 and nucleosides and nucleotides19 with derivatization.The sensitivity of photothermal absorption techniques is 2–3 orders of magnitude higher than those of conventional detection methods. Since the absorption wavelength of analytes is required to match the emission wavelength of the pump beam well and the sensitivity of photothermal detection is related to the molar absorptivity of the analytes, these detectors are therefore not suitable for detecting non-absorbing analytes. This is one important reason why crossed-beam photothermal (CBPT) techniques have not been employed in CE for determinations of metal ions.In this paper, an indirect photothermal interference detection for detecting metal cations without derivatization after CE separation is described. Methylene Blue was chosen as a background absorber. The addition of ethanol and sodium chloride to the background electrolyte solution (BGES) can reduce the absorption of Methylene Blue and metal cations on the capillary wall, enhance the detector signal and improve the separation efficiency.Experimental Apparatus The indirect photothermal interference detector is shown in Fig. 1. A 2 mW He–Ne laser (Wuhan Institute of Laser Technology, Wuhan, China) produces a probe beam with a wavelength of 632.8 nm. A 46 mm focal length biconvex lens is used to focus the probe beam into the capillary tube. After traversing the capillary tube, the beam propagates to a photodiode (Semiconductor Factory, Wuhan University, Wuhan, China) with a 0.5 mm slit width.An M1000 He–Ne laser (Shanghai Institute of Laser Technology, Shanghai, China) produces an 18 mW pump laser beam. A model 194A mechanical chopper (EG&G, Princeton Applied Research, Princeton, NJ, USA) modulates the pump beam in a square wave. This beam is focused with a 10 mm focal length biconvex lens into the capillary tube. A three-axis translation stage holds the pump beam lens to provide complete freedom in the location of the pump beam waist, a two-axis stage is used to move the capillary, which is fixed firm on a special installation with a pinhole along the beam path of the probe laser, in the plane formed by the two laser beams, and a two-axis stage moves the detector across the probe beam profile.The interaction of the pump laser beam with the samples results in a refractive index change within the sample. This refractive index change causes a movement of the interference fringes and a change in the probe beam intensity which is synchronized with the pump beam modulation function. The modulated component of the probe beam intensity is measured by using an ND240 lock-in amplifier (Nanjing University Instrument Factory, Nanjing, China). A reference signal for the lock-in amplifier is generated within the chopper head.During the process of the electrophoresis, the signals from the lock-in amplifier are recorded by our SE 790 recorder (Australia). The capillary electrophoresis system was constructed in the laboratory.A high-voltage dc power supply (0–30 kV) (Shanghai, China) is employed. Electrical connections to the capillary are enclosed within a plastic box equipped with a safety interlock that acts to prevent accidental contact with the high voltage. A 60 cm long polymide-coated fused-silica capillary (Yongnian Optical Fiber Factory, Hebei, China) of 50 mm id and 320 mm od is used for electrophoresis. The polymer coating of the detection region is scraped away carefully.The distance from the injector to the detector is 50 cm. During the experiment, the optical breadboard is held at ground potential. Fig. 1 Schematic diagram of the detection system. PL, probe laser; M1 and M2, mirrors; L1 and L2, lenses; Ph1 and Ph2, pinholes; Ch, chopper; C, capillary; HL, heat laser; D, photodiode; LI, lock-in amplifier; R, recorder. 1090 Analyst, October 1997, Vol. 122Reagents All reagents were of analytical-reagent grade and solutions were prepared by dissolution in doubly distilled, de-ionized water.Oxides of metals were obtained from Shanghai Chemical (Shanghai, China). Samples were diluted by using the carrier electrolyte to avoid stacking and introduced into the capillary by using hydrostatic injection from a height of 10 cm. BGES were diluted with the same carrier electrolyte. All solutions and the buffer were filtered with 0.22 mm cut-off cellulose acetate filters before use. This filtration greatly reduces noise spikes and prevents blockage of the small capillaries.All data were obtained at room temperature (25 ± 1 °C). Results and Discussion Choice of Background Absorber The characteristics of the background absorber are very important for detection and separation with the utilization of indirect absorption detection in CZE. A higher molar absorptivity (ev) at the detection wavelength and a low CM will give a lower detection limit, as described by Wang and Hartwick.20 Five color reagents were obtained in order to assess their suitability as indirect background absorbers before separation.Experiments showed that the absorption wavelengths of pyrocatechol and Eriochrome Black T could match the pump laser beam (632.8 nm). However, these color reagents were unstable during the separation process and faded easily, because phenolic hydroxyl radicals are easily oxidized in an air environment. Victoria Blue B and Coomassie Brilliant Blue G 250 are not easily dissolved in aqueous solution.The solubility could be improved by adding ethanol, acetone or dimethylformamide to the electrolyte solution, but it was found that some small solid grains appeared on evaporation of the solvent, and these blocked the capillaries. Therefore, the color reagents mentioned above are not suitable for detection or separation in CZE. The molar absorptivity of Methylene Blue at 633 nm is about 4.5 3 104 l mol21 cm21. Experiments showed that characteristics of Methylene Blue are suitable for indirect interference detection and CZE separation.Noise Analysis It was found that the noise in indirect photothermal interference detection includes at least three sources: the detector, the color reagent and Joule heating. A significant effort was made to minimize the vibration of optical components. For example, the optical components are held with massive fixtures to a wall-damped optical table. When a laser beam strikes a capillary, a scattered light beam with many light and dark fringes can be observed, as described by Folestad et al.21 and Kerker and Matijewc22 and this scattered light beam is an important noise source in detection.First, portions of this scattered light, coming from the scattered light beam produced by the probe laser and the pump laser, could retroreflect back to both the probe laser source and pump beam laser source. This kind of retroreflection could cause fluctuations of the laser source and increase the noise level.Second, it was found that the noise level increases when the portion of scattered light coming from the scattered light beam produced by the pump laser hits the photodiode in the far-field after the capillary. This part of the scattered light is modulated by a mechanical chopper, and both the pump laser beam and probe laser beam are at the same wavelength in this system. In the present work, the probe laser beam was focused into the capillary at a right-angle, which tilted up to the capillary tube axis (a Å 89°). After being focused by a biconvex lens, the modulated pump laser beam was tilted slightly down into the capillary tube axis (b Å 89°).Further, the pump laser beam and the probe beam crossed obliquely at the capillary at a rightangle (g Å 89°) and two pinholes were used to extinguish the scattered light from other optical components, as shown in Fig. 1. It was demonstrated that the noise caused by the scattered light could be minimized successfully in this configuration. The reason is that the beams of scattered light in the plane of the interaction of the laser beam and the capillary were tilted to the capillary tube axis and not easy to retroreflect to the laser source and photodiode.As noted by Higashijima et al.23 the use of Methylene Blue in an acidic buffer leads to peak tailing characteristic of absorption of the positively charged dye on the capillary surface and would lead to higher noise.As shown later in Fig. 4, this type of absorption on the capillary surface could be minimized by adding ethanol to the electrolyte solutions and using an electrolyte solution of higher ionic strength. This phenomenon can be attributed to the fact that ethanol and a higher ionic strength can limit the dissociation of silanol on the capillary surface and the absorption of Methylene Blue. Further, ethanol and a higher ionic strength also enhanced the Methylene Blue detection signal, because the dn/dT of solutions containing ethanol and a higher ionic strength electrolyte is larger than that of aqueous solutions.24 This was beneficial for improving the sensitivity of indirect photothermal interference detection.It was also found that an excessively low concentration of the color reagent would lead to higher noise, as the ratio of the background absorbance to the noise somehow depends on the concentration of the color reagent and decreases simultaneously. 25 Excessive Joule heating should be avoided in CE as it prevents efficient heat dissipation to attain high efficiencies and causes thermal instability in the detection region.In general, as described in the Introduction, a low ionic strength of the buffers is required in indirect detection. However, an important effect was found in our experiments, namely that, in a certain range, an increase in the separation voltage in CZE did not decrease but actually improved the sensitivity of indirect photothermal interference detection. The reason is that the temperature rise in the detection region inside the capillary would maintain a stable level under the conditions of a stable voltage,26 and this kind of temperature increase of the background electrolyte could lead to an increase in the dn/dT value, as demonstrated by Franko and Tran.27 The contribution to noise from the Joule heating was found to be small under the condition of a stable voltage.Indirect Detection Before detection for CZE separation, several alignment constraints were qualitatively investigated for photothermal capillary absorption measurements. As expected, the signal was highest when the pump laser beam waist was centered in the capillary. Since a tightly focused pump beam also minimizes the system volume, all measurements reported in this paper were obtained with the pump waist located in the capillary. For a given interference pattern, the fringes that were found to be more sensitive were those that appear near the optical axis of the probe beam but still retain high intensity and contrast.When the interference pattern and the slit width of detector were fixed, an optimum distance for detection from the sample cuvette to the photodiode was observed. Agreeing well with a lower noise level and higher sensitivity, in the experiment the photodiode was set at a position of 60 cm in the far-field after the capillary.Under these conditions, the capillary was located at a position of 19 mm before the probe beam waist to obtain higher sensitivity. When the capillary containing the solutions was drawn gradually closer to the probe beam waist, it was found that the signal decreased quickly. The reason may be that the contrast of the interference fringes also decreased quickly. Analyst, October 1997, Vol. 122 1091A finite time is required to reach thermal equilibrium in photothermal absorbance measurements. As a general rule of thumb, a frequency of 6.5 Hz was used in these experiments. A short time constant is of value since very fast phenomena may be studied without degradation due to instrumental artifacts, and the detector time constant was set at 1 s.An absorbance calibration curve of Methylene Blue was obtained in the static state for the 50 mm id capillaries. The calibration curve was linear (r = 0.995) over more than three orders of magnitude of concentration. Linearity extended from the detection limit, A = 6.1 31027, to the highest concentration sample studied, A = 0.023.Water–ethanol solution was chosen as the solvent because it produces greater sensitivity than water for photothermal absorbance measurements. As shown in Fig. 2, for Methylene Blue solution, the concentration LOD was about 3.1 3 1028 mol l21 (S/N = 2). With a highly collimated beam, the detection volume of this detector, the intersection volume of the pump and probe beam, is small, slightly less than 1 pl.We use a much more conservative definition of the detection volume, as described by Bornhop and Dovichi,28 as a cylinder whose radius and height are given by the capillary radius. For the 50 mm id tube, the detector volume was about 50 pl. Then, the detection limit as absolute amount was 1.5 3 10218 mol. Of course, a significantly smaller amount of analyte was present within the small intersection volume of the laser beam than the result estimated above.It has been demonstrated that, for absorption measurement in narrow-bore capillaries, crossed-beam photothermal (CBPT) absorption techniques can offer definite advantages: the sensitivities are not linear with respect to pathlength, and with a decrease in the capillary diameter the pump beam is more efficiently quenched at the walls. Therefore, according to the assumption described in ref. 28, this detector could also be used with a capillary tube as small as 5 mm id. The results in Fig. 3 show that the addition of ethanol to the BGES can reduce absorption on the capillary surface and lead to an improvement in the peak shape of the analyte. Ethanol also enhanced the Methylene Blue and analyte detection signal, because the dn/dT value of ethanolic solutions is larger than that of aqueous solutions, and this is beneficial for the improvement of sensitivity in indirect interference detection.Further, it could be observed that the elution time became longer with an increase of ethanol concentration. A probable cause is that the ethanol interacts strongly with the capillary wall, resulting in a higher apparent concentration of ethanol within the double layer as described in ref. 29. These interactions then result in higher apparent viscosities within the double layer and lead to a reduced electroosmotic flow. The effect of NaCl on CZE separation and detection was investigated.The results (Fig. 4) showed that efficiency of KI and the peak shape were improved at higher concentrations of NaCl without a decrease in detection sensitivity. These results and that of the Joule heating phenomenon (see above) indicated that indirect photothermal interference detection is suitable for CE separation systems with a high ionic strength buffer. As the sensitivity that CBPT provides was almost unrelated to the light path, this result implies that indirect photothermal interference detection may be used for detection in CZE with a larger ionic strength range and overcomes the problem of the decrease in sensitivity for small id capillaries and shot-noise limitation in photometry.The characteristics of photothermal absorbance detection and the results obtained above are, for example, beneficial for the analysis of samples with weak absorption, ultratrace detection and larger range of linearity, and permit a lower concentration of the background absorber (Methylene Blue) to be used to improve the LOD.As shown in Fig. 5, positive peaks are observed for KI and CuII and AlIII produces a negative peak. A positive peak indicates an increase in concentration of the background absorber present at the detector, whereas a negative peak indicates a decrease. The migration time of the system peak corresponds to the mobility of the Methylene Blue cation. An interesting phenomenon is observed in Fig. 5, namely that the migration order is KI > CuII > AlIII. In order to understand the elution order, it has to be taken into account that the electrophoretic mobility depends on the charge, shape and size of the analyte. This phenomenon could be explained by the fact that, under conditions of low pH and excess of chloride, Fig. 2 Electropherogram of Methylene Blue. Carrier electrolyte, 50 mmol l21 Na2HPO4–10% ethanol (pH 6.0); sample, 5 3 1027 mol l21 Methylene Blue; CE voltage, 16 kV; current, 14 mA; injection, 10 s.Fig. 3 Electropherograms of KI (1.0 3 1025 mol l21). Carrier electrolyte; (a) 109 mmol l21 acetic acid–5% ethanol; (b) 109 mmol l21 acetic acid 210% ethanol; and (c) 109 mmol l21 acetic acid–20% ethanol. CE voltage, 14 kV; injection, 15 s; Methylene Blue concentration, 5.0 3 1024 mol l21. Fig. 4 Electropherogram of KI (1.0 3 1025 mol l21). Carrier electrolyte, 109 mmol l21 acetic acid–20% ethanol–100 mmol l21 NaCl; CE voltage, 14 kV; current, 24 mA; other conditions as in Fig. 3. 1092 Analyst, October 1997, Vol. 122CuCl4 22 and AlCl42 complexes are formed in solution, but KI is incapable of forming chloride complexes. Therefore, the peak of KI appeared first. The elution order of CuCl4 22 and AlCl42 can be rationalized by the fact that AlCl42 seems to be a poorly hydrated anion compared with CuCl4 22. Hence this lower hydration will reduce the size of the moving ion, which may result in greater electrophoretic migration and, accordingly, retardation. The phenomenon is similar to the studies of Aguilar et al.30 and Baraj et al.,31 in which chloride and cyanide were used as ligands for the determination of metal cations.For CuII, the samples and carrier solution were prepared with almost the same conductivity, and consequently the stacking effect was not used; a 2.1 3 1027 mol l21 concentration detection limit (S/N = 2) without preconcentration and a 1.20 3 10217 mol detection limit as absolute amount, considering the optical sampling volume estimated to be 50 pl, were measured, and 1.99 3106 theoretical plates were observed with the laboratorymade CE system. The concentration LODs measured here were comparable to the best results obtained with other detection methods for ions,32–34 and the mass LOD of this detector could be decreased to that of an indirect fluorescence detector or conductivity detector (10217 mol) and at least two orders of magnitude lower than that of a commercial UV detector.The CE method offers several advantages over other indirect detection methods for ions. First the use of a larger range of ionic strengths permits the application dynamic range to be increased35,36 and extends the upper limit of sample ionic strength that still permits effective stacking and thus facilitates the determination of trace constituents in a high ionic strength matrix. Second, the lower mass LOD permits small id capillaries to be used to improve the resolution of samples and ultra trace components. Obviously, this characteristic of indirect photothermal interference detection is better than that of commercial detection.Even then the best mass LODs of conductimetric detection and indirect fluorescence detection are similar to those of this method; as one mode of absorption detection techniques, this method has all the advantages of photometric detection over conductimetric detection mentioned earlier, and maybe a less noisy baseline than that in indirect fluorescence detection, since the main disadvantage of indirect fluorescence detection is that fluorescence gives rise to a noisier baseline than does absorption and leads to a higher concentration LOD.Third, as the values of dn/dT for most non-aqueous media are larger than those of aqueous solutions, this detector will be of advantage for detection in non-aqueous medium capillary electrophoresis, an important field in the study of capillary electrophoresis.In future work we will investigate possible improvements to the detection sensitivity, including the design of an electronic noise canceller to reduce electronic noise, altering the injection mode and the use of highly stable lasers. The relatively low cost, simple construction, small volume, universal detection, larger range of ionic strengths and excellent detection limits of this method should prove attractive for a number of applications in capillary electrophoresis separation techniques. This work was supported by the National Nature Science Foundation of China. References 1 Stoker, F. S., Haymor, B. L., and McBeath, R., J. Chromatogr., 1989, 470, 241. 2 Burlity, N., and Jorgenson, J., J. Chromatogr., 1989, 480, 301. 3 Swaile, D. F., and Sepaniak, M. J., Anal. Chem., 1991, 63, 179. 4 Hu, Y., Deng, Y., and Cheng, J., Prog. Nat. Sci., 1996, 6, S-42. 5 Yeung, E. S., Acc. Chem. Res., 1989, 22, 125. 6 Yeung, E. S., and Kuhr, W. G., Anal. Chem., 1991, 63, 275A. 7 Kaniansky, D., Zelenska, V., and Baluchova, D., Electroporesis, 1996, 17, 1890. 8 Jandik, P., and Bonn, G., Capillary Electrophoresis of Small Ions, VCH, Weinheim, 1993. 9 Pfeffer, W. D., and Yeung, E. S., J. Chromatogr., 1990, 506, 401. 10 Wilson, S. A., and Yeung, E. S., Anal. Chim. Acta, 1984, 157, 53. 11 Mikkers, F. E. P., Everaerts, F. M., and Verheggen, Th. P. E. M., J. Chromatogr., 1979, 169, 11. 12 Kuhr, W. G., and Yeung, E. S., Anal. Chem., 1988, 60, 2642. 13 Takeuchi, T., and Yeung, E. S., J. Chromatogr., 1986, 370, 83. 14 Gussman, E., Kuo, R. N., and Zare, R. N., Science, 1985, 230, 813. 15 Pawliszyn, J., J. Liq. Chromatogr., 1987, 10, 3377. 16 Cheng, C. Y., and Morris, M. D., Appl. Spectrosc., 1988, 42, 515. 17 Yu, M., and Dovich, N. J., Mikrochimica III, 1988, 27. 18 Yu, M., and Dovich, N. J., Anal. Chem., 1989, 61, 37. 19 Krattiger, B., Bruno, A. E., Widmer, H. M., and Pandliker, R., Anal. Chem., 1995, 67, 124. 20 Wang, T. S., and Hartwick, R. A., J. Chromatogr., 1992, 607, 119. 21 Folestad, S., Johnson, L., Josefsson, B., and Galle, B., Anal. Chem., 1982, 54, 925. 22 Kerker, M., and Matijewc, E., J. Opt. Soc. Am., 1961, 51, 506. 23 Higashijima, T., Fuchigami, T., Imasaka, T., and Ishibashi, N., Anal. Chem., 1992, 64, 711. 24 Fujiwara, K., Lei, W., Uckiki, H., Shimokoshi, F., Fuwa, K., and Kobayashi, T., Anal. Chem., 1982, 54, 2026. 25 Takenchi, T., and Yeung, E. S., J. Chromatogr., 1986, 370, 83. 26 Bruno, A. E., Krattiger, B., Maystre, F., and Widmer, H. M., Anal. Chem., 1991, 63, 2689. 27 Franko, M., and Tran, C. D., J. Phys. Chem., 1991, 95, 6688. 28 Bornhop, D. J., and Dovichi, N. J., Anal. Chem., 1987, 59, 1632. 29 VanOrman, B. B., Liversidge, G. G., and McIntire, G. L., J. Microcolumn Sep., 1991, 2, 176. 30 Aguilar, M., Farran, A., and Martinnez, M., J. Chromatogr., 1993, 635, 127. 31 Baraj, B., Sastre, A., Merkoci, A., and Martinnez, M., J. Chromatogr., 1995, 718, 227. 32 Weston, A., Broun, P. R., Jandik, P., Heckenberg, A. L., and Jones, W. R., J. Chromatogr., 1992, 608, 395. 33 Kuhr, W. G., and Yeung, E. S., Anal. Chem., 1988, 60, 2642. 34 Kaniansky, D., Zelenska, V., and Baluchova, D., Electrophoresis, 1996, 17, 1890. 35 Nielen, M. W. F., J. Chromatogr., 1991, 542, 173. 36 Green, J. S., and Jorgenson, J. W., J. Chromatogr., 1987, 478, 63. Paper 7/00119C Received January 6, 1997 Accepted June 18, 1997 Fig. 5 Electropherogram of a mixture of three metal cations. Methylene Blue concentration, 5.0 3 1026 mol l21. Other conditions as in Fig. 4. Peaks: 1, KI (2.5 3 1026 mol l21); 2, system peak; 3, CuII (1.56 3 1026 mol l21); and 4, AlIII (3.85 3 1026 mol l21). Analyst, October 1997, Vol. 122 1093

ISSN:0003-2654    

DOI:10.1039/a700119c

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

CH2 N S H H COOH O CONH The Effect of Cooking on Veterinary Drug Residues in Food. Part 8. Benzylpenicillin† Martin D. Rose*, John Bygrave, William H. H. Farrington and George Shearer CSL Food Science Laboratory, Norwich Research Park, Colney, Norwich, UK NR4 7UQ The stability of benzylpenicillin to heat and cooking was studied. Stability of this compound in water (at 100 °C and 65 °C), 5% ethanol, 5% sodium bicarbonate, pH 5.5 buffer at 100 °C and in hot cooking oil at 140 °C and 180 °C was established.Benzylpenicillin was stable at 65 °C but not stable at higher temperatures with half-life times varying between 15 and 60 min in the solutions investigated. This drug was not stable to cooking, losses being proportional to the harshness of the cooking regime. Where fluids were released during the cooking process, sometimes over half of the residue passed from the solid tissue into the cooking medium. Keywords: Benzylpenicillin; thermal stability; cooking; veterinary drug residues; antibiotic; food Benzylpenicillin (penicillin G) is a widely used antibiotic in both veterinary and human medicine.The structure of this compound is shown in Fig. 1. Concern has been shown over the possible presence of residues of this drug in foods of animal origin due to the occurrence of penicillin hypersensitivity in humans. In veterinary medicine it may be used both to treat specific infections and also as a prophylactic. It is widely used by the dairy industry for the treatment of mastitis in cows.Benzylpenicillin is administered as one or more of a variety of salts which are used to prolong the activity of the drug. These can be the soluble sodium or potassium salts or the longer acting procaine and benzathine salts. The maximum residue limit (MRL) for benzylpenicillin is 4 mg kg21 in milk and 50 mg kg21 in any other edible tissue.1 Both the chemical stability of benzylpenicillin and the microbiological condition of the individual food item needs to be addressed when considering the effects of storage and cooking on this residue.The presence of micro-organisms which produce the enzyme penicillinase could be capable of reducing the concentration of penicillin in food during storage and cooking.2 It has been suggested that observed decreases in the incidence of benzylpenicillin residues in milk in recent years may be attributable to the addition of this enzyme or enzyme producing bacteria to the milk.2 It is of interest that benzylpenicillin is reported to be more stable in UHT treated milk than in raw milk.3 A few studies on the effect of storage and heat treatment of benzylpenicillin residues in food have been carried out.4–7 Most of these4–6 used a microbiological screening assay but one study which used TLC and autoradiochromatography identified degradation products.8 In this study small samples of freeze dried meat soaked in a spiking solution were used.Analysis was carried out on liquid squeezed from the cooked or stored samples. The study did not produce fully quantitative data since only a portion of the radioactivity was recovered. Some decrease in antimicrobial activity was seen for storage of kidney tissue at 4 °C. Cooking could also result in some decrease in activity, although significant activity remained from procaine penicillin with ‘rare’ and ‘medium’ cooking conditions in hamburgers, steaks and pork chops.Only a small percentage of original activity was left after ‘well done’ or ‘exceptionally well done’ cooking. It was shown that the major breakdown product formed during both storage and cooking was the lactate ester of penicilloic acid.8 Antimicrobial activity was reduced in the production of different sausage types with a suggestion that inactivation was due to penicillinase-producing microbes.9 The aim of this study was to use HPLC methodology to produce qualitative and quantitative data about the change in concentration of benzylpenicillin residues in food cooked by a variety of methods.Experimental Benzylpenicillin Heat Stability Experiments to determine heat stability in water were performed at 100 °C and 65 °C. Glass crimp-top autosampler vials (2 ml capacity) were filled with 10 mg ml21 benzylpenicillin solution in water and placed in either a 1 l beaker of boiling water on a hot plate for measurement of stability at 100 °C or in a beaker of water previously equilibrated in a thermostatically controlled air-circulating oven at 65 °C.Samples were removed at intervals, cooled rapidly and analysed by HPLC together with controls of unheated benzylpenicillin solution. To determine heat stability in 5% ethanol, vials were filled with 10 mg ml21 benzylpenicillin in 5% ethanol solution to simulate cooking food with wine or beer. They were placed in a thermostatically controlled air-circulating oven at 100 °C. Samples were removed at intervals and treated as above.To determine heat stability in 5% sodium bicarbonate solution, vials were filled with 10 mg ml21 benzylpenicillin in 5% sodium bicarbonate solution (pH 8.2) and placed in a beaker of boiling water to investigate stability at 100 °C under mildly basic cooking conditions. To determine heat stability in acidic buffer, vials were filled with 10 mg ml21 benzylpenicillin solution in 0.15 m phosphate buffer (pH 5.5) and placed in a beaker of boiling water to investigate stability under acidic conditions.This pH was chosen because it approximates to that of meat.10 Two investigations of heat stability in hot cooking oil were carried out. The first was at 140 °C and the second at 180 °C, a temperature typically used for domestic cooking. Sunflower oil (200 g) was placed in a beaker containing a glass-coated magnetic stirring rod and heated using an electric hot plate. † For Part 7, see ref. 12. Fig. 1 Structure of benzylpenicillin.Analyst, October 1997, Vol. 122 (1095–1099) 1095Temperature was monitored using a mercury in glass thermometer. Benzylpenicillin solution (200 ml) at 1 mg ml21 in methanol was added to give a concentration of 1 mg g21 benzylpenicillin in oil. Sub-samples of at least 5 g were removed at intervals over a period of 2 h. These were placed in 20 ml test tubes which were cooled rapidly by immersion in cold water. Exactly 5 g of oil was mixed with 5 ml hexane and extracted into 5 ml 0.01 m phosphate buffer (pH 8.5).Benzylpenicillin was measured by HPLC together with control standards, prepared by extracting unheated oil spiked at an equivalent concentration. The temperatures achieved during each cooking procedure were monitored using a fibre thermometer. The model used was suitable for temperature measurement between 0–200 °C with an accuracy of 0.1 °C. Samples Benzylpenicillin incurred cattle muscle was obtained from the BBSRC Institute for Animal Health, Compton, Berkshire, UK.The cattle liver was taken from an animal of weight 310 kg dosed by injection with 8.2 ml solution containing 300 mg ml21 procaine benzylpenicillin (Norocillin, Norbrook Laboratories, Kidderminster, UK). This animal was slaughtered 48 h after dosing. The muscle tissue from this animal contained insufficient residues of penicillin to measure. Homogenised muscle tissue from the injection site of a different animal treated for another project was used.This was not an ideal matrix to represent consumer use but since minced beef muscle is used in many standard recipes and this was a source of tissue with a measurable amount of incurred residues it was preferable to the use of tissue fortified with penicillin post mortem. The animal from which this tissue was obtained weighed 100 kg and was treated with a double dose (2 injection sites, left and right rump) of 4 ml solution containing 112.5 mg ml21 benzathine benzylpenicillin and 150 mg ml21 procaine benzylpenicillin.Tissue from these animals was stored at 270 °C until required for use since it has been shown that residues of penicillin are not stable at conventional freezer temperatures. Blank tissues for method validation and quality control samples were obtained from local retail outlets. They were analysed prior to use using the method described below under Analytical Method to verify the absence of benzylpenicillin residues. Cooking Procedure A single piece of tissue was used for each cooking investigation. A portion of raw tissue was removed and analysed in duplicate or triplicate and the remainder was cooked as a single sample.Replicate sub-samples of the cooked tissue were analysed for residues of benzylpenicillin. Samples of water used, or exudate formed during cooking were also analysed. Exact composition of the number of replicates of each sample type was dependent upon the availability of sample, and analytical batch size.All raw incurred samples of a particular tissue type used in the cooking investigations were sub-samples originating from the same animal. Cooking Methods Investigated A variety of cooking methods were used to reflect the wide variation in domestic practise. Specific recipes were as described below. Cattle liver and muscle by frying A slice of cattle liver (156 g) was fried with a little sunflower oil in a pan using an electric hob.The liver was cooked for 20 min, turning occasionally. The cooked liver had a ‘well done’ appearance on the outside but was still pink in the centre. A small amount of homogenised cattle muscle (29 g) was fried in a little sunflower oil on low heat from a gas flame for 10 min. Cattle muscle by boiling, microwave and roasting Water and salt were added to homogenised cattle muscle (73 g) and the mixture was brought to the boil. The mixture was simmered for 13 min. Homogenised cattle muscle (31 g) was cooked for 3 min without the addition of water in a casserole dish with glass lid in a 700 W microwave oven set on medium power.Homogenised cattle muscle (84 g) was cooked in a casserole dish with glass lid in a gas oven, mark 6 (205 °C) for 30 min. No water was added. Cattle liver by braising and casserole Cattle liver (59 g) was fried in a little sunflower oil over a gas flame at low heat for 10 min. Water was added and the mixture brought to the boil and simmered for an additional 10 min.Liver (66 g) was placed in a casserole dish with 60 ml salted water. The sample was cooked at gas mark 7 (220 °C) for 28 min. Analytical Method The analytical method used was based on that developed by Boison et al.11 The method used penicillin V as internal standard for quantification and recovery measurements. This was used at concentrations of 1 mg kg21 for liver experiments and 10 mg kg21 for muscle, to reflect the concentration of the residue in each tissue type.Benzylpenicillin was extracted from tissue with acetone and cleaned up using C18 solid phase extraction (SPE). The eluate from the SPE cartridge was derivatized with 1,2,4-triazole–mercuric chloride and the derivative was subject to reversed-phase HPLC. A schematic diagram of the procedure is shown in Fig. 2. Batches consisted normally of between three and six samples and two blank tissues, one fortified with benzylpenicillin at an appropriate concentration in addition to internal standard.The fortified extract was used as a recovery. A calibration curve was constructed using a range of benzylpenicillin standards prepared in mobile phase covering the concentrations found in the incurred tissue extracts. Results and Discussion Method Validation The method used for the analysis of residues of benzylpenicillin was validated for raw and cooked tissue before use. The validation data obtained for the method are included in Table 1.The method has been previously validated in this laboratory with tissue fortified with 10 mg kg21 benzylpenicillin and the limit of detection is estimated at below 5 mg kg21 which agrees with the figure given by Boison et al. in the original description of the method.11 The method is in routine use in our laboratory and had previously been demonstrated to give a linear response for the range of concentrations measured in this investigation. Heat Stability of Benzylpenicillin Plots of benzylpenicillin concentration against time at 100 °C in various aqueous systems are shown in Fig. 3. At 65 °C benzylpenicillin was stable in water as shown in Fig. 4. The half-life time at 100 °C in water and 5% ethanol was approximately 60 min. This reduced to about 40 min in acidic buffer (pH 5.5) and about 15 min in 5% sodium bicarbonate solution (pH 8.2) at the same temperature. Plots of benzylpen- 1096 Analyst, October 1997, Vol. 122icillin concentration against time in oil at 140 °C and 180 °C are shown in Fig. 5; the approximate half-life times were 45 min and 20 min respectively. This suggested that benzylpenicillin was less stable in an aqueous than in a lipid environment, and indicated that degradation was caused by acid or base catalysed hydrolysis. It was previously established that the likely products from heating benzylpenicillin in simple solution are benzylpenillic acid, isomers of benzylpenicilloic acid, isomers of benzylpenilloic acid and penicillenic acid.8 Effect of Cooking on Residue Concentration The results for the effect of cooking on benzylpenicillin residues in food are all corrected for recovery and are shown in Table 2.The amount of penicillin in food decreased as it was cooked, except for homogenised cattle muscle, by boiling for 10 min. No significant changes in concentration were observed for boiling, when compared with the analytical precision of the method. The boiling experiment was repeated to verify this result because it was not consistent with the results from the other cooking methods.The findings were, however, consistent with the prediction based on the model obtained from heating benzylpenicillin standard in water for the same amount of time. The losses seen for other methods were dependent on cooking time and conditions, the greatest losses observed were for processes which used the highest temperatures and longest times; these were homogenised cattle muscle by frying where a 90% loss was found and cattle liver by braising where a 75% loss was found.The maximum temperature in the frying experiment was higher than any of the other processes at 160 °C for the exterior surfaces and the cooking time was 10 min for muscle and 20 min for liver. Braising involved frying the cattle liver for 10 min where the exterior surface reached a fairly high temperature (up to about 120 °C) followed by simmering for 10 min where the sample was cooked at a constant 100 °C.Variations between replicate analyses for liver were greater than for homogenised muscle probably because of the temperature variation between the surface and interior of the sample. For the cattle liver by casserole experiment a loss of 79% is reported. The concentration in raw tissue was averaged from duplicate measurements of 0.94 and 0.45 mg kg21. The range of the loss Fig. 2 Schematic of analytical procedure.Table 1 Method validation data. Recovery (%) from tissue fortified with benzylpenicillin Cattle muscle Cattle liver Raw Cooked Raw Cooked Day 1/ Day 2/ Day 3/ Day 1/ Day 1/ Day 1/ Sample No. 100 mg kg21 100 mg kg21 100 mg kg21 10 mg kg21 200 mg kg21 1 mg kg21 1 45.2 52.0 59.1 54.9 58.5 46.0 2 49.2 54.2 63.6 72.1 61.8 49.2 3 39.3 52.1 63.3 57.7 58.2 64.7 4 50.8 53.8 55.2 64.8 60.1 54.2 5 38.9 50.9 60.8 61.8 6 65.8 54.4 72.4 Mean (%) 44.7 54.8 59.4 64.0 59.8 53.5 sn21 5.49 5.53 3.94 7.27 1.7 8.2 RSD (%) 12.3 10.1 6.63 11.4 2.9 15.2 n 5 6 6 6 4 4 Fig. 3 Benzylpenicillin stability at 100 °C in water, 5% ethanol, 5% sodium bicarbonate and pH 5.5 buffer. Analyst, October 1997, Vol. 122 1097was between 67% and 85% based on these two starting concentrations. The analytical precision can be assessed from the validation data and the difference between replicate analysis of the homogenised raw muscle tissue. The mean loss was in line with predictions that may have been made on the basis of data obtained from the model systems where drug was heated in simple solutions.The variation in recovery for different cooking methods reflected day to day performance of the method as seen when it is applied to the surveillance of raw tissue samples. The benzylpenicillin which was measured after cooking was found to be distributed between the solid cooked tissue and liquids which were used for cooking or which came from the meat as it was cooked.Sometimes (for casserole and boiling homogenised cattle muscle) over half of the total measured residue was present in the fluid. Distribution of Residues Within the Incurred Tissue No assessment of the distribution of residues within the muscle tissue was possible. This muscle tissue was homogenised prior to cooking, and results from the analysis of raw samples associated with each cooking experiment showed that there was a small decrease in the starting concentration of benzylpenicillin in raw tissue during the course of the work.Samples were kept at 270 °C for long term storage, but were kept at 220 °C for the Fig. 4 Benzylpenicillin stability in water at 65 °C. Table 2 Effect of cooking on benzylpenicillin residues in food Homogenised cattle muscle Cattle liver Fried Microwave Casserole Boiled Boiled Fried Braised Casserole Sample No. 10 min 3 min 30 min 13 min (1) 13 min (2) 20 min 20 min 28 min Recovery (%) 76 76 80 87 100 54 66 51 Raw— 1 13.5 13.5 11.9 12.5 9.9 0.33 0.81 0.94 2 16.2 16.2 10.7 14.8 9.7 0.23 0.82 0.45 3 0.37 Mean/mg kg21 14.9 14.9 11.3 13.7 9.8 0.31 0.82 0.69 Cooked— 1 3.1 8.1 1.9 7.1 4.5 0.25 0.09 0.05 2 2.8 8.3 1.9 6.9 4.7 0.23 0.23 0.13 3 2.7 6.9 2.3 8.1 4.8 0.34 0.08 4 1.8 6.6 4.9 0.07 0.05 5 7.8 0.26 0.04 6 0.22 0.15 Mean/mg kg21 2.8 7.8 2.0 7.3 4.7 0.24 0.20 0.08 RSD (%) 7.2 10.2 11.6 8.6 3.4 51.3 55.8 Fluid— 1 Insufficient Insufficient 1.40 9.22 3.2 Insufficient 0.08 0.08 2 sample sample 1.50 9.22 3.1 sample 0.07 0.10 3 1.41 9.36 3.2 0.10 4 1.40 3.2 0.09 Mean/mg kg21 1.43 9.27 3.2 0.07 0.09 Mass/g— Raw 29.1 31.3 84.4 72.8 50.0 156 58.9 66.1 Cooked 15.0 24.3 94.5 74.5 48.1 129 49.5 50.4 Fluid 0 0 177 90.1 94.0 0 28.1 55.6 Total benzylpenicillin/mg— Raw 434 466 954 997 490 48 48 46 Cooked 43 189 186 544 228 31 9.9 4.5 Fluid — — 253 415 297 — 2 5.2 Net change (%) 290 259 254 24 +7 236 275 279 Fig. 5 Benzylpenicillin stability in cooking oil at 140 °C and 180 °C. 1098 Analyst, October 1997, Vol. 122duration of most of the experimental work which took place over a relatively short (2–3 week) time period. This elevation in temperature was sufficient to affect the stabilty of penicillin residues. It is possible that the process of homogenising raw tissue could have an effect on benzylpenicillin. It was nevertheless preferable to use this tissue which originated from a treated animal than to use only tissue fortified by the addition of drug prior to analysis. Results from experiments carried out on this tissue were in line with results for the liver experiments and with the findings from the experiments conducted on model systems.The analytical precision for cooked liver was much poorer than that expected for analytical error alone and may reflect the difference in temperature between the internal and external parts of the sample during cooking. Conclusions Benzylpenicillin was found to be unstable during cooking in these experiments which were performed on muscle tissue from one animal and liver from a second.Much of the residue migrated with juices which exuded from the tissue as it was cooked. No major metabolites or breakdown products were monitored or identified in this study. The findings supported the losses found using different methodology in a previous investigation where the lactate ester of penicilloic acid was identified as the major product formed in cooked food.8 Surveillance data obtained for measurements of benzylpenicillin in raw tissue may not therefore be directly applicable for use in consumer exposure estimates and dietary intake calculations for the cooked product.Exposure to residues of benzylpenicillin may be reduced by discarding any juices which come from the meat as it is cooked. References 1 EEC, 1992. Council Regulation (EEC) No 675/92 Amending Annexes I and III of Council Regulation (EEC) No 2377/90 laying down a Community procedure for the establishment of maximum residue limits of veterinary medicinal products in foodstuffs of animal origin.(OJ No L 73, 19.3.1992, pp. 8–14). 2 Guay, R., Cardinal, P., Bourassa, C., and Brassard, N., Int. J. Food Microbiol., 1987, 4, 187. 3 Haagsma, N., in Proceedings of the Euroresidue II Conference on Residues of Veterinary Drugs in Food, Veldhoven, Netherlands, 3–5 May 1993, ed. Haagsma, N., Ruiter, A., and Czedic-Eysenberg, P. B., pp. 41–49. 4 Katz, S. E., Fassbender, C.A., DePaolis, A. M., and Rosen, J. D., J. Assoc. Off. Anal. Chem., 1978, 61(3), 564. 5 O’Brien, J. J., Campbell, N., and Conaghan, T., J. Hyg., 1981, 87, 511. 6 Pilet, Ch., and Toma, B., Etude sur la thermostabilit�e de quelques antibiotiques. Rec. Med. Vet., 1969, 145(9), 897. 7 Boison, J. O., Korsrud, G. O., MacNeil, J. D., and Yates, W. D. G., J. AOAC Int., 1992, 75(6), 974. 8 DePaolis, A. M., Katz, S. E., and Rosen, J. D., J. Agric. Food Chem., 1978, 25(5), 1112. 9 Scheibner,G., Monatsh. Veterinaermed., 1972, 27, 161. 10 Price, J. F, and Schweigert, B. S., The science of meat and meat products, Freeman, San Francisco, 2nd edn., 1971. 11 Boison, J. O. K., Salisbury, C. D.C., Chan, W., and MacNeil, J. D., J. AOAC Int., 1991, 74, 497. 12 Rose, M., Farrington, W., and Shearer, G., Food Addit. Contam., in the press. Paper 7/02771K Received April 23, 1997 Accepted July 16, 1997 Analyst, October 1997, Vol. 122 1099 CH2 N S H H COOH O CONH The Effect of Cooking on Veterinary Drug Residues in Food.Part 8. Benzylpenicillin† Martin D. Rose*, John Bygrave, William H. H. Farrington and George Shearer CSL Food Science Laboratory, Norwich Research Park, Colney, Norwich, UK NR4 7UQ The stability of benzylpenicillin to heat and cooking was studied. Stability of this compound in water (at 100 °C and 65 °C), 5% ethanol, 5% sodium bicarbonate, pH 5.5 buffer at 100 °C and in hot cooking oil at 140 °C and 180 °C was established.Benzylpenicillin was stable at 65 °C but not stable at higher temperatures with half-life times varying between 15 and 60 min in the solutions investigated. This drug was not stable to cooking, losses being proportional to the harshness of the cooking regime. Where fluids were released during the cooking process, sometimes over half of the residue passed from the solid tissue into the cooking medium. Keywords: Benzylpenicillin; thermal stability; cooking; veterinary drug residues; antibiotic; food Benzylpenicillin (penicillin G) is a widely used antibiotic in both veterinary and human medicine.The structure of this compound is shown in Fig. 1. Concern has been shown over the possible presence of residues of this drug in foods of animal origin due to the occurrence of penicillin hypersensitivity in humans. In veterinary medicine it may be used both to treat specific infections and also as a prophylactic. It is widely used by the dairy industry for the treatment of mastitis in cows. Benzylpenicillin is administered as one or more of a variety of salts which are used to prolong the activity of the drug.These can be the soluble sodium or potassium salts or the longer acting procaine and benzathine salts. The maximum residue limit (MRL) for benzylpenicillin is 4 mg kg21 in milk and 50 mg kg21 in any other edible tissue.1 Both the chemical stability of benzylpenicillin and the microbiological condition of the individual food item needs to be addressed when considering the effects of storage and cooking on this residue. The presence of micro-organisms which produce the enzyme penicillinase could be capable of reducing the concentration of penicillin in food during storage and cooking.2 It has been suggested that observed decreases in the incidence of benzylpenicillin residues in milk in recent years may be attributable to the addition of this enzyme or enzyme producing bacteria to the milk.2 It is of interest that benzylpenicillin is reported to be more stable in UHT treated milk than in raw milk.3 A few studies on the effect of storage and heat treatment of benzylpenicillin residues in food have en carried out.4–7 Most of these4–6 used a microbiological screening assay but one study which used TLC and autoradiochromatography identified degradation products.8 In this study small samples of freeze dried meat soaked in a spiking solution were used.Analysis was carried out on liquid squeezed from the cooked or stored samples.The study did not produce fully quantitative data since only a portion of the radioactivity was recovered. Some decrease in antimicrobial activity was seen for storage of kidney tissue at 4 °C. Cooking could also result in some decrease in activity, although significant activity remained from procaine penicillin with ‘rare’ and ‘medium’ cooking conditions in hamburgers, steaks and pork chops. Only a small percentage of original activity was left after ‘well done’ or ‘exceptionally well done’ cooking.It was shown that the major breakdown product formed during both storage and cooking was the lactate ester of penicilloic acid.8 Antimicrobial activity was reduced in the production of different sausage types with a suggestion that inactivation was due to penicillinase-producing microbes.9 The aim of this study was to use HPLC methodology to produce qualitative and quantitative data about the change in concentration of benzylpenicillin residues in food cooked by a variety of methods.Experimental Benzylpenicillin Heat Stability Experiments to determine heat stability in water were performed at 100 °C and 65 °C. Glass crimp-top autosampler vials (2 ml capacity) were filled with 10 mg ml21 benzylpenicillin solution in water and placed in either a 1 l beaker of boiling water on a hot plate for measurement of stability at 100 °C or in a beaker of water previously equilibrated in a thermostatically controlled air-circulating oven at 65 °C.Samples were removed at intervals, cooled rapidly and analysed by HPLC together with controls of unheated benzylpenicillin solution. To determine heat stability in 5% ethanol, vials were filled with 10 mg ml21 benzylpenicillin in 5% ethanol solution to simulate cooking food with wine or beer. They were placed in a thermostatically controlled air-circulating oven at 100 °C. Samples were removed at intervals and treated as above. To determine heat stability in 5% sodium bicarbonate solution, vials were filled with 10 mg ml21 benzylpenicillin in 5% sodium bicarbonate solution (pH 8.2) and placed in a beaker of boiling water to investigate stability at 100 °C under mildly basic cooking conditions.To determine heat stability in acidic buffer, vials were filled with 10 mg ml21 benzylpenicillin solution in 0.15 m phosphate buffer (pH 5.5) and placed in a beaker of boiling water to investigate stability under acidic conditions.This pH was chosen because it approximates to that of meat.10 Two investigations of heat stability in hot cooking oil were carried out. The first was at 140 °C and the second at 180 °C, a temperature typically used for domestic cooking. Sunflower oil (200 g) was placed in a beaker containing a glass-coated magnetic stirring rod and heated using an electric hot plate. † For Part 7, see ref. 12. Fig. 1 Structure of benzylpenicillin. Analyst, October 1997, Vol. 122 (1095–1099) 1095Temperature was monitored using a mercury in glass thermometer. Benzylpenicillin solution (200 ml) at 1 mg ml21 in methanol was added to give a concentration of 1 mg g21 benzylpenicillin in oil. Sub-samples of at least 5 g were removed at intervals over a period of 2 h. These were placed in 20 ml test tubes which were cooled rapidly by immersion in cold water. Exactly 5 g of oil was mixed with 5 ml hexane and extracted into 5 ml 0.01 m phosphate buffer (pH 8.5).Benzylpenicillin was measured by HPLC together with control standards, prepared by extracting unheated oil spiked at an equivalent concentration. The temperatures achieved during each cooking procedure were monitored using a fibre thermometer. The model used was suitable for temperature measurement between 0–200 °C with an accuracy of 0.1 °C. Samples Benzylpenicillin incurred cattle muscle was obtained from the BBSRC Institute for Animal Health, Compton, Berkshire, UK.The cattle liver was taken from an animal of weight 310 kg dosed by injection with 8.2 ml solution containing 300 mg ml21 procaine benzylpenicillin (Norocillin, Norbrook Laboratories, Kidderminster, UK). This animal was slaughtered 48 h after dosing. The muscle tissue from this animal contained insufficient residues of penicillin to measure. Homogenised muscle tissue from the injection site of a different animal treated for another project was used.This was not an ideal matrix to represent consumer use but since minced beef muscle is used in many standard recipes and this was a source of tissue with a measurable amount of incurred residues it was preferable to the use of tissue fortified with penicillin post mortem. The animal from which this tissue was obtained weighed 100 kg and was treated with a double dose (2 injection sites, left and right rump) of 4 ml solution containing 112.5 mg ml21 benzathine benzylpenicillin and 150 mg ml21 procaine benzylpenicillin.Tissue from these animals was stored at 270 °C until required for use since it has been shown that residues of penicillin are not stable at conventional freezer temperatures. Blank tissues for method validation and quality control samples were obtained from local retail outlets. They were analysed prior to use using the method described below under Analytical Method to verify the absence of benzylpenicillin residues. Cooking Procedure A single piece of tissue was used for each cooking investigation.A portion of raw tissue was removed and analysed in duplicate or triplicate and the remainder was cooked as a single sample. Replicate sub-samples of the cooked tissue were analysed for residues of benzylpenicillin. Samples of water used, or exudate formed during cooking were also analysed. Exact composition of the number of replicates of each sample type was dependent upon the availability of sample, and analytical batch size.All raw incurred samples of a particular tissue type used in the cooking investigations were sub-samples originating from the same animal. Cooking Methods Investigated A variety of cooking methods were used to reflect the wide variation in domestic practise. Specific recipes were as described below. Cattle liver and muscle by frying A slice of cattle liver (156 g) was fried with a little sunflower oil in a pan using an electric hob. The liver was cooked for 20 min, turning occasionally.The cooked liver had a ‘well done’ appearance on the outside but was still pink in the centre. A small amount of homogenised cattle muscle (29 g) was fried in a little sunflower oil on low heat from a gas flame for 10 min. Cattle muscle by boiling, microwave and roasting Water and salt were added to homogenised cattle muscle (73 g) and the mixture was brought to the boil. The mixture was simmered for 13 min. Homogenised cattle muscle (31 g) was cooked for 3 min without the addition of water in a casserole dish with glass lid in a 700 W microwave oven set on medium power.Homogenised cattle muscle (84 g) was cooked in a casserole dish with glass lid in a gas oven, mark 6 (205 °C) for 30 min. No water was added. Cattle liver by braising and casserole Cattle liver (59 g) was fried in a little sunflower oil over a gas flame at low heat for 10 min. Water was added and the mixture brought to the boil and simmered for an additional 10 min.Liver (66 g) was placed in a casserole dish with 60 ml salted water. The sample was cooked at gas mark 7 (220 °C) for 28 min. Analytical Method The analytical method used was based on that developed by Boison et al.11 The method used penicillin V as internal standard for quantification and recovery measurements. This was used at concentrations of 1 mg kg21 for liver experiments and 10 mg kg21 for muscle, to reflect the concentration of the residue in each tissue type.Benzylpenicillin was extracted from tissue with acetone and cleaned up using C18 solid phase extraction (SPE). The eluate from the SPE cartridge was derivatized with 1,2,4-triazole–mercuric chloride and the derivative was subject to reversed-phase HPLC. A schematic diagram of the procedure is shown in Fig. 2. Batches consisted normally of between three and six samples and two blank tissues, one fortified with benzylpenicillin at an appropriate concentration in addition to internal standard.The fortified extract was used as a recovery. A calibration curve was constructed using a range of benzylpenicillin standards prepared in mobile phase covering the concentrations found in the incurred tissue extracts. Results and Discussion Method Validation The method used for the analysis of residues of benzylpenicillin was validated for raw and cooked tissue before use. The validation data obtained for the method are included in Table 1. The method has been previously validated in this laboratory with tissue fortified with 10 mg kg21 benzylpenicillin and the limit of detection is estimated at below 5 mg kg21 which agrees with the figure given by Boison et al.in the original description of the method.11 The method is in routine use in our laboratory and had previously been demonstrated to give a linear response for the range of concentrations measured in this investigation. Heat Stability of Benzylpenicillin Plots of benzylpenicillin concentration against time at 100 °C in various aqueous systems are shown in Fig. 3. At 65 °C benzylpenicillin was stable in water as shown in Fig. 4. The half-life time at 100 °C in water and 5% ethanol was approximately 60 min. This reduced to about 40 min in acidic buffer (pH 5.5) and about 15 min in 5% sodium bicarbonate solution (pH 8.2) at the same temperature. Plots of benzylpen- 1096 Analyst, October 1997, Vol. 122icillin concentration against time in oil at 140 °C and 180 °C are shown in Fig. 5; the approximate half-life times were 45 min and 20 min respectively.This suggested that benzylpenicillin was less stable in an aqueous than in a lipid environment, and indicated that degradation was caused by acid or base catalysed hydrolysis. It was previously established that the likely products from heating benzylpenicillin in simple solution are benzylpenillic acid, isomers of benzylpenicilloic acid, isomers of benzylpenilloic acid and penicillenic acid.8 Effect of Cooking on Residue Concentration The results for the effect of cooking on benzylpenicillin residues in food are all corrected for recovery and are shown in Table 2.The amount of penicillin in food decreased as it was cooked, except for homogenised cattle muscle, by boiling for 10 min. No significant changes in concentration were observed for boiling, when compared with the analytical precision of the method. The boiling experiment was repeated to verify this result because it was not consistent with the results from the other cooking methods.The findings were, however, consistent with the prediction based on the model obtained from heating benzylpenicillin standard in water for the same amount of time. The losses seen for other methods were dependent on cooking time and conditions, the greatest losses observed were for processes which used the highest temperatures and longest times; these were homogenised cattle muscle by frying where a 90% loss was found and cattle liver by braising where a 75% loss was found.The maximum temperature in the frying experiment was higher than any of the other processes at 160 °C for the exterior surfaces and the cooking time was 10 min for muscle and 20 min for liver. Braising involved frying the cattle liver for 10 min where the exterior surface reached a fairly high temperature (up to about 120 °C) followed by simmering for 10 min where the sample was cooked at a constant 100 °C.Variations between replicate analyses for liver were greater than for homogenised muscle probably because of the temperature variation between the surface and interior of the sample. For the cattle liver by casserole experiment a loss of 79% is reported. The concentration in raw tissue was averaged from duplicate measurements of 0.94 and 0.45 mg kg21. The range of the loss Fig. 2 Schematic of analytical procedure. Table 1 Method validation data.Recovery (%) from tissue fortified with benzylpenicillin Cattle muscle Cattle liver Raw Cooked Raw Cooked Day 1/ Day 2/ Day 3/ Day 1/ Day 1/ Day 1/ Sample No. 100 mg kg21 100 mg kg21 100 mg kg21 10 mg kg21 200 mg kg21 1 mg kg21 1 45.2 52.0 59.1 54.9 58.5 46.0 2 49.2 54.2 63.6 72.1 61.8 49.2 3 39.3 52.1 63.3 57.7 58.2 64.7 4 50.8 53.8 55.2 64.8 60.1 54.2 5 38.9 50.9 60.8 61.8 6 65.8 54.4 72.4 Mean (%) 44.7 54.8 59.4 64.0 59.8 53.5 sn21 5.49 5.53 3.94 7.27 1.7 8.2 RSD (%) 12.3 10.1 6.63 11.4 2.9 15.2 n 5 6 6 6 4 4 Fig. 3 Benzylpenicillin stability at 100 °C in water, 5% ethanol, 5% sodium bicarbonate and pH 5.5 buffer. Analyst, October 1997, Vol. 122 1097was between 67% and 85% based on these two starting concentrations. The analytical precision can be assessed from the validation data and the difference between replicate analysis of the homogenised raw muscle tissue. The mean loss was in line with predictions that may have been made on the basis of data obtained from the model systems where drug was heated in simple solutions.The variation in recovery for different cooking methods reflected day to day performance of the method as seen when it is applied to the surveillance of raw tissue samples. The benzylpenicillin which was measured after cooking was found to be distributed between the solid cooked tissue and liquids which were used for cooking or which came from the meat as it was cooked. Sometimes (for casserole and boiling homogenised cattle muscle) over half of the total measured residue was present in the fluid.Distribution of Residues Within the Incurred Tissue No assessment of the distribution of residues within the muscle tissue was possible. This muscle tissue was homogenised prior to cooking, and results from the analysis of raw samples associated with each cooking experiment showed that there was a small decrease in the starting concentration of benzylpenicillin in raw tissue during the course of the work.Samples were kept at 270 °C for long term storage, but were kept at 220 °C for the Fig. 4 Benzylpenicillin stability in water at 65 °C. Table 2 Effect of cooking on benzylpenicillin residues in food Homogenised cattle muscle Cattle liver Fried Microwave Casserole Boiled Boiled Fried Braised Casserole Sample No. 10 min 3 min 30 min 13 min (1) 13 min (2) 20 min 20 min 28 min Recovery (%) 76 76 80 87 100 54 66 51 Raw— 1 13.5 13.5 11.9 12.5 9.9 0.33 0.81 0.94 2 16.2 16.2 10.7 14.8 9.7 0.23 0.82 0.45 3 0.37 Mean/mg kg21 14.9 14.9 11.3 13.7 9.8 0.31 0.82 0.69 Cooked— 1 3.1 8.1 1.9 7.1 4.5 0.25 0.09 0.05 2 2.8 8.3 1.9 6.9 4.7 0.23 0.23 0.13 3 2.7 6.9 2.3 8.1 4.8 0.34 0.08 4 1.8 6.6 4.9 0.07 0.05 5 7.8 0.26 0.04 6 0.22 0.15 Mean/mg kg21 2.8 7.8 2.0 7.3 4.7 0.24 0.20 0.08 RSD (%) 7.2 10.2 11.6 8.6 3.4 51.3 55.8 Fluid— 1 Insufficient Insufficient 1.40 9.22 3.2 Insufficient 0.08 0.08 2 sample sample 1.50 9.22 3.1 sample 0.07 0.10 3 1.41 9.36 3.2 0.10 4 1.40 3.2 0.09 Mean/mg kg21 1.43 9.27 3.2 0.07 0.09 Mass/g— Raw 29.1 31.3 84.4 72.8 50.0 156 58.9 66.1 Cooked 15.0 24.3 94.5 74.5 48.1 129 49.5 50.4 Fluid 0 0 177 90.1 94.0 0 28.1 55.6 Total benzylpenicillin/mg— Raw 434 466 954 997 490 48 48 46 Cooked 43 189 186 544 228 31 9.9 4.5 Fluid — — 253 415 297 — 2 5.2 Net change (%) 290 259 254 24 +7 236 275 279 Fig. 5 Benzylpenicillin stability in cooking oil at 140 °C and 180 °C. 1098 Analyst, October 1997, Vol. 122duration of most of the experimental work which took place over a relatively short (2–3 week) time period. This elevation in temperature was sufficient to affect the stabilty of penicillin residues. It is possible that the process of homogenising raw tissue could have an effect on benzylpenicillin. It was nevertheless preferable to use this tissue which originated from a treated animal than to use only tissue fortified by the addition of drug prior to analysis. Results from experiments carried out on this tissue were in line with results for the liver experiments and with the findings from the experiments conducted on model systems.The analytical precision for cooked liver was much poorer than that expected for analytical error alone and may reflect the difference in temperature between the internal and external parts of the sample during cooking. Conclusions Benzylpenicillin was found to be unstable during cooking in these experiments which were performed on muscle tissue from one animal and liver from a second.Much of the residue migrated with juices which exuded from the tissue as it was cooked. No major metabolites or breakdown products were monitored or identified in this study. The findings supported the losses found using different methodology in a previous investigation where the lactate ester of penicilloic acid was identified as the major product formed in cooked food.8 Surveillance data obtained for measurements of benzylpenicillin in raw tissue may not therefore be directly applicable for use in consumer exposure estimates and dietary intake calculations for the cooked product. Exposure to residues of benzylpenicillin may be reduced by discarding any juices which come from the meat as it is cooked. References 1 EEC, 1992. Council Regulation (EEC) No 675/92 Amending Annexes I and III of Council Regulation (EEC) No 2377/90 laying down a Community procedure for the establishment of maximum residue limits of veterinary medicinal products in foodstuffs of animal origin. (OJ No L 73, 19.3.1992, pp. 8–14). 2 Guay, R., Cardinal, P., Bourassa, C., and Brassard, N., Int. J. Food Microbiol., 1987, 4, 187. 3 Haagsma, N., in Proceedings of the Euroresidue II Conference on Residues of Veterinary Drugs in Food, Veldhoven, Netherlands, 3–5 May 1993, ed. Haagsma, N., Ruiter, A., and Czedic-Eysenberg, P. B., pp. 41–49. 4 Katz, S. E., Fassbender, C. A., DePaolis, A. M., and Rosen, J. D., J. Assoc. Off. Anal. Chem., 1978, 61(3), 564. 5 O’Brien, J. J., Campbell, N., and Conaghan, T., J. Hyg., 1981, 87, 511. 6 Pilet, Ch., and Toma, B., Etude sur la thermostabilit�e de quelques antibiotiques. Rec. Med. Vet., 1969, 145(9), 897. 7 Boison, J. O., Korsrud, G. O., MacNeil, J. D., and Yates, W. D. G., J. AOAC Int., 1992, 75(6), 974. 8 DePaolis, A. M., Katz, S. E., and Rosen, J. D., J. Agric. Food Chem., 1978, 25(5), 1112. 9 Scheibner,G., Monatsh. Veterinaermed., 1972, 27, 161. 10 Price, J. F, and Schweigert, B. S., The science of meat and meat products, Freeman, San Francisco, 2nd edn., 1971. 11 Boison, J. O. K., Salisbury, C. D.C., Chan, W., and MacNeil, J. D., J. AOAC Int., 1991, 74, 497. 12 Rose, M., Farrington, W., and Shearer, G., Food Addit. Contam., in the press. Paper 7/02771K Received April 23, 1997 Accepted July 16, 1997 Analyst, October 1997, Vol. 122 10

ISSN:0003-2654    

DOI:10.1039/a702771k

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Bioelectrochemical Determination of Citric Acid in Real Samples Using a Fully Automated Flow Injection Manifold Mamas I. Prodromidisa, Stella M. Tzouwara-Karayannia, Miltiades I. Karayannis*a and Pankaj M. Vadgamab a University of Ioannina, Laboratory of Analytical Chemistry, Ioannina 45110, Greece b University of Manchester, Department of Medicine, Hope Hospital, Salford, UK M6 HD An enzymic method for the determination of citric acid in fruits, juices and sport drinks is proposed.The method is based on the action of the enzymes citrate lyase, oxaloacetate decarboxylase and pyruvate oxidase, which convert citric acid into H2O2 with the latter being monitored amperometrically with a H2O2 probe. The enzymes pyruvate oxidase and oxaloacetate decarboxylase were immobilized. A multi-membrane system, consisting of a cellulose acetate membrane for the elimination of interferants, an enzymic membrane and a protective polycarbonate membrane were placed on a Pt electrode and used with a fully automated flow injection manifold.Several parameters were optimized, resulting in a readily constructed and reproducible biosensor. Interference from various compounds present in real samples was minimized. Calibration graphs were linear over the range 0.01–0.9 mm pyruvate, 0.015–0.6 mm oxaloacetate and 0.015–0.5 mm citrate. The throughput was 30 samples h21 with an RSD of 1.0% (n = 8); the mean relative error was 2.4% compared with a standard method.The recovery was 96–104%. A 8–10% loss of the initial activity of the sensor was observed after 100–120 injections. Keywords: Amperometric citrate determination; multimembrane biosensor; flow injection analysis; fruits, juices and sport drinks Citric acid is present in numerous natural products and is a key tribasic acid involved in both plant and animal aerobic respiration. Several fresh fruits such as lemons and limes owe their sharp taste to the presence of the citrate anion.1 Citric acid is also an additive in industry, mainly as a preservative and an acidulant. It is added to some dairy products to improve and protect both flavor and aroma.It has been used for radioactive Sr2+ from suspect milk after radiation fallout, and has been useful for chelating trace metals which can cause haze or deterioration of color and flavor.2 Several methods have been proposed for the determination of citric acid, based on ion-exchange chromatography,3 HPLC4 and isotachophoresis;5 these are time consuming procedures, and sample clean-up is required in order to separate citric acid from other co-existing tricarboxylic acids.Other approaches based on conductimetric6 or spectrophotometric methods7 suffer from selectivity as they are based on non-specific reactions with carboxylic acids. The need for simple separation could be potentially solved, by using a high specificity enzyme such as citrate lyase8 (CL). Moellering and Gruber9 reported a method with soluble CL, lactate dehydrogenase and malate dehydrogenase (MDH), and monitored the decrease in the absorbance of the NADH at 340 nm.Although relatively slow and expensive in enzyme the approach works well, as a batch method, and is widely used in routine off-line analysis. Since then several other approaches have been proposed; some make use of soluble CL and immobilized MDH with NADH10 detection, some use CL and oxaloacetate decarboxylase (OACD) in soluble11 or immobilized12 form in conjunction with polarography.The direct amperometric determination of citric acid has been proposed13 in connection with an ascorbate oxidase reactor to eliminate ascorbic acid interference. Gajovic et al.14 used CL, purified by recrystallisation and ultrafiltration and then entrapped in gelatin. This method needs recalibration every six runs probably owing to the gradual loss of the activity of the CL. The enzyme could be reactivated after 1 h incubation with adenosine 5A- triphosphate (ATP) and CH3COOH.Measurements using a non-enzymic membrane were a proposed route to the elimination of interferent species. CL is usually used in soluble form because of ready inactivation by complex formation with Mg2+ and Zn2+ and also the enol-form of oxaloacetate,15 itself the product of its action on citrate. The proposed method is based on a sequence of reactions involving thiamine pyrophosphate (TPP) and the enzymes CL, OACD and pyruvate oxidase (POD), according to the following scheme: citrate oxaloacetate CH COOH CL 3 ¾ ®¾ + (1) oxaloacetate pyruvate + CO OACD 2 ¾ ® ¾¾ (2) pyruvate + HPO O acetyl-phosphate CO H O 4 2 2 2 2 POD TPP, Mg 2+ 2-+ ® + + (3) The final product H2O2 was monitored amperometrically by means of a Pt electrode, mounted on a wall-jet flow-through cell, polarised at +0.65 V versus an Ag–AgCl reference electrode. The multi-membrane biosensor gave an interferantfree system, with good analytical characteristics in terms of accuracy, reproducibility and operational simplicity, which is a key advantage of biosensors over conventional enzymic assays.Additionally, the system was combined with a flow injection (FI) manifold, fully automated by means of resident reported software.16 Experimental Apparatus This work was carried out using an in-house fully automated FI manifold. Electrochemical experiments were run using a computer-controlled potentiostat (Eco Chemie/Autolab, Utrecht, The Netherlands).Solutions were pumped through the manifold using a four-channel peristaltic pump (Gilson, Villiers Analyst, October 1997, Vol. 122 (1101–1106) 1101le Bel, France) and sample injections made with a pneumatically actuated injection valve (Rheodyne, Cotati, CA, USA). A resident program, synchronized with the acquisition software, ensured full control of pump and valves, eliminating manual manipulation; a detailed description of the program is given elsewhere.16 A three-electrode flow-through detector (Metrohm 656, Herisau, Switzerland) was used for the amperometric monitoring.This consists of a wall-jet thermostated cell (volume < 1 ml) with a Pt (diameter 1.6 mm, Bioanalytical Systems, W. Lafayette, IN, USA) working electrode, a built-in Au auxiliary electrode and an Ag–AgCl reference electrode. A schematic diagram of the system is shown in Fig. 1(A). Chemicals POD (EC 1.2.3.3, from Pediococcus sp., 43 U mg21), OACD (EC 4.1.1.3, from Pseudommonas sp., 265 U mg21), CL (EC 4.1.3.6, from Enterobacter aerogenes, 0.26 U mg21) all in lyophilized form and cellulose acetate (approximately 40% acetyl), were obtained from Sigma (St.Louis, MO, USA). 3- Morpholinopropanesulfonic acid (MOPS) and poly(vinyl acetate) (PVA, Mr 167 000 Da) were supplied by Aldrich (Gollingham, Germany). The enzymic test kit for citric acid determinations was purchased from Boehringer (Mannheim, Germany).All other (analytical-reagent grade) chemicals were purchased from Sigma. Solutions Working (MOPS 70 mm, pH 7.6) and immobilization (MOPS 50 mm, pH 7.35) buffer solutions were filtered through a 1.0 mm pore diameter microporous polycarbonate (PC) membrane (Millipore, Bedford, MA, USA), prior to use. Stock solutions of pyruvic (50 mm) and oxaloacetic acid (50 mm) were prepared by dissolving 27.5 and 33.1 mg of sodium pyruvate and oxaloacetic acid in 5.0 ml of 0.1 and 0.5 m HCl, respectively; this prevented their polymerization or decarboxylation.Both solutions were stored at 4 °C and prepared weekly, owing to their instability.17 A stock solution of TPP (20 mm, pH 6.2) was prepared by dissolving 181.4 mg of TPP in 20 ml of the working buffer. Enzyme solutions were prepared in the immobilization buffer, otherwise working buffer solution was used throughout. Membranes Polymeric nylon 66 membranes, thickness 120 mm, viz., Biodyne A (amphoteric, 50% amino, 50% carboxyl, 0.2 mm porosity), Biodyne B (positively charged with pore surfaces populated by a high density of quaternary ammonium groups, 0.45 mm porosity), Biodyne C (negatively charged, contains 100% carboxyl, 0.45 mm porosity) and Immunodyne ABC (preactivated, 0.2 mm porosity) were a kind gift from Pall Filtration (Milan, Italy).HA (mixed ester cellulose, 0.45 mm porosity) membranes were purchased from Millipore. PC membranes (thickness 10 mm, 0.03 and 0.05 mm porosity) and dialysis tubing (12 000 Mr cut-off) were supplied by Nuclepore (Cambridge, MA, USA) and Sigma, respectively.For casting cellulose acetate membranes a wet film applicator (5 in, 1–8 mils, URAI, Milan, Italy) was used. Preparation of the Enzymic Membranes POD was immobilized by ionic and/or covalent bonding onto nylon 66 membranes, following the spot wetting method:18 A 10 ml (10 U) aliquot of the POD solution was symmetrically pipetted onto each side of the dry membrane (diameter 6 mm) and left to air-dry for 5 min. Alternatively, physical entrapment of POD was carried out by passing 50 ml (10 U) of POD solution through an HA membrane (diameter 6 mm) while slight suction was applied.19 In both cases, unattached protein was removed by washing the membranes (3 3 10 min) with the immobilization buffer.POD and OACD were co-immobilized asymmetrically onto Biodyne B and Immunodyne ABC membranes, according to a procedure reported in the literature:20 A 10 ml (10 U) aliquot of the POD solution was applied onto the rough side of the dry membrane (1 3 1 cm2) and allowed to react for 2–3 min.Then, 6 ml (15 U) of the OACD solution were applied onto the smooth side of the membrane and allowed again to react for 15 min. The membranes were washed and stored in the immobilization buffer at +4 °C. Assembly of the Sensor The cellulose acetate membrane (20 mm thick, 100 Da cut-off) was first placed on the Pt surface to eliminate interference from electroactive species.21 This membrane was prepared in our laboratory by dissolving 3.996 g of cellulose acetate and 4 mg of PVA in a mixture of 60 ml of acetone and 40 ml of cyclohexanone (after a minor modification of the procedure reported by Palleschi et al.21).Then the membrane bearing the enzyme(s) was superimposed with an outer PC membrane in order to prevent microbial attack and also leaching of enzyme. With the oxaloacetate biosensor, the rough surface of the membrane was directed towards the cellulose acetate; POD membranes could be placed with any orientation.All the Fig. 1 (A) Schematic representation of FI manifold employed for citric acid determinations: C, carrier; R, TPP reagent. (B) Assembly of the biosensor; LM, large molecules; AN, analyte; IN, interferants; Pt, platinum electrode. CA, cellulose acetate; EM, enzymic membrane; PC, polycarbonate membrane. (C) i, POD membrane; ii, random application; and iii, asymmetric application. 1102 Analyst, October 1997, Vol. 122membranes were tightly fitted over the electrode with the aid of an O-ring [Fig. 1(B)]. PC membranes of different porosity (0.03 and 0.05 mm) and a dialysis membrane (modified with 1% Na2CO3 and 10 mm Trizma–1 mm EDTA22) were alternatively used as protective membranes. After preliminary evaluation of the effect of these membranes on the pyruvate biosensor, in terms of response times, sensitivity, linearity and operational stability, a porosity of 0.03 mm was finally selected. Preparation of Sample Solutions A 5 g amount of fruit sample was homogenized with a blender, diluted in 100 ml of the working buffer and finally filtered through a 1.0 mm PC membrane.Juice and sport drink samples were directly diluted in 10 ml of working buffer, while carbonated samples were sonicated (1 min, 100 W) prior to dilution. Procedure The carrier (70 mm MOPS of pH 7.6 containing 3.22 nm MgCl2 and 1.61 mm KH2PO4) and TPP [2.64 mm, pH 6.2 (2.64 ml of 20 mm TPP in 20 ml of water)] streams were continuously pumped at flow rates of 0.23 and 0.14 ml min21, respectively, towards the probe until a stable baseline current was reached (1–2 nA, within 10–15 min).Standard or sample solutions of citric acid [40 ml (0.4 U) CL + x ml sample + (960 2x) ml carrier] were introduced as short pulses of 120 ml via the loop injection valve. For the oxaloacetate or pyruvate biosensor, the composition of the standards was [x ml standard + (5000 2 x) ml carrier].The peak height of the current response was taken as a measure of the analyte concentration. Results and Discussion Enzyme Membranes Commencing from the first enzymic step [eqn. (3)], POD was immobilized onto several commercially available membranes. The wide range of the chemical and physical properties of the membranes led us to find the most efficient matrix for the construction of the pyruvate biosensor.As well as the membrane spot wetting immobilization, membrane immersion18 was tested using a 0.32 mg per 2 ml (10 U POD) enzymic solution; the latter gave poor immobilization efficiencies. Enzyme loading was tested separately for POD and OACD and defined the enzyme loading necessary to obtain diffusional limitation of response, i.e., the response maximum.14 Saturation study of the membrane was made using the Immunodyne ABC membrane; this is claimed to have the greatest binding capacity. The response to pyruvate with different enzyme loading solutions (2–12 U POD) was studied (Fig. 2). Experiments were then performed, applying the useful enzyme content (10 U POD) onto different membranes. The relative apparent measurable efficiencies, which reflect the biocatalytic efficiency of the immobilized enzymes, are shown in Table 1. Biodyne B and Immunodyne ABC membranes showed the best results and so further work was carried out using these membranes.By keeping the amount of POD at the optimum value, further membranes were prepared by varying the amount of the applied OACD from 5 to 20 U. The responses of oxaloacetate biosensors with different enzyme loadings are shown in Fig. 2. Asymmetric co-immobilization of POD and OACD gave greater current signals than those with random application of the same amount of the enzymes [Fig. 1(C)]. Probe Lifetime POD is a relatively unstable enzyme probably owing to the gradual loss of its cofactor flavin adenine dinucleotide (FAD), during both the immobilization procedure and the measurements.The addition of several compounds such as FAD, TPP, MgCl2, glycerol and (NH4)2SO4 to the immobilization or storage buffer did not improve either the immobilization efficiency or the long-term stability. Indeed, this led to a more complicated storage of the membranes and greater cost. OACD is a more stable enzyme; therefore, lifetime experiments were carried out with the pyruvate biosensor (Table 2).The proposed approach is more economical than other attempts, with similar13 or slightly better lifetimes23 where the preparation time of the enzymic membranes and the amount of the enzymes are significantly greater ( > 30 h preparation time and 10–26-fold amounts of the enzyme). Concentration of Activators and Cofactors It has been reported24 that the activity of POD is affected by the presence of inorganic phosphate, divalent cations such as Mg2+, Ca2+, Mn2+ or Co2+ and cofactors such as TPP and FAD.This optimization was carried out with the probe assembled as a pyruvate biosensor, using 0.5 mm pyruvate solution as a sample. Fig. 2 Enzyme loading test using the Immunodyne ABC membrane: 5 POD (2–12 U); -, POD (10 U) + OACD (5–20 U); 0.5 mm substrate. Parameters: 70 mm MOPS; pH 7.5; 1 mm MgCl2; 2 mm TPP; 1 mm KH2PO4. Table 1 Relative efficiencies of various polymeric membranes used as supports for the immobilization of POD. 0.3 mm pyruvate; 10 U POD Membrane Current/nA RE (%)* Biodyne A 29.3 68 Biodyne B 42.8 100 Biodyne C 27.3 64 Immunodyne ABC 40.7 95 Millipore HA 22.1 52 * Relative efficiency with respect to the most active membrane. Table 2 Lifetime of the pyruvate biosensor. Immunodyne ABC membrane; 10 U POD; 0.2 mm pyruvate* Day Current/nA Activity (%) 1 26.0 100 2 23.4 90 3 20.7 80 4 19.0 73 5 9.2 35 6 5.5 21 7 1.0 4 * Continuous use: 100–200 injections (8–10% loss of the initial activity).Use after storage: minimum 15 injections per day. Analyst, October 1997, Vol. 122 1103TPP was initially included in the carrier stream, but it was found to be unstable in this environment (decay of signal output), and so was pumped separately [Fig. 1(A)] achieving, furthermore, better mixing with the sample solution. In the absence of TPP no response was observed while for concentrations higher than 0.8 mm TPP the response became constant; subsequent work used 1mM TPP.Several cations were examined as activators of POD. In addition to sensitivity, the criterion for the final selection was the stability of the sensor. Since Ca2+ and Co2+ precipitate with phosphate they must be avoided; Ca2+ also competed with the binding site for Mg2+, vital for CL activity. A 2 mm MgCl2 concentration was found to be optimum for the performance of the biosensor. Inorganic phosphate, as KH2PO4, was tested in the concentration range 0–2 mm and a concentration of 1 mm was selected; the graph was similar to that seen with Mg2+.FAD in the carrier stream (up to 0.5 mm) had no effect on the current output; it is likely that the enzyme preparation contains sufficient FAD for the catalytic reaction and no further FAD was needed. Using POD–OACD membranes and 0.15 mm oxaloacetate, the effect of TPP, MgCl2 and KH2PO4 was re-examined and similar optima were observed. The effect of Mn was further investigated and it was found to increase the activity of the system (approximately 6–8%) when a concentration of 0.1 mm MnCl2 was added to the carrier stream.At higher concentrations a progressive decrease of the signal was observed, accompanied by the formation of Mn3(PO4)2; hence, subsequent work was carried out in the absence of MnCl2. Working Conditions The pH of the working buffer was also investigated in several buffering systems, such as MOPS, glycylglycine and Trizma- HCl, covering the pH range 7–8.The last gave lower responses, probably owing to its reaction with Mg2+ and oxaloacetate ions. Using POD–OACD membranes, sequential injections of 0.3 mm pyruvate and 0.3 mm oxaloacetate, at different pH values of 70 mm MOPS, were performed. The optimum pH for both the pyruvate and oxaloacetate biosensors is 7.5 as shown in Fig. 3. The numbers that appear on the graph were calculated as (S1/S2) 3 100 (where S1 and S2 represent the signals obtained for standards of 0.3 mm oxaloacetate and pyruvate, respectively) and reflect the efficiency of the conversion of oxaloacetate to pyruvate. The efficiencies of the conversion were also investigated at different flow rates (reaction times), following the procedure given above.Flow rate profiles are shown in Fig. 4. An overall flow rate of 0.37 ml min21 was finally selected, which reconciles fairly high peaks and satisfactory sample throughput (30 h21). A sample volume of 120 ml was used as it prevented peak broadening (dispersion coefficient 1.22–1.25) and also ensured high sensitivity.The sensitivity of the oxaloacetate biosensor also increased with the temperature, levelling off at a maximum value of 47 °C. Above this temperature thermal inactivation dominates over the increase in the collision frequency, resulting in a decrease of the signal. All experiments were carried out at 30 °C, where the stability of the biosensor was the same as at room temperature. Amount of Citrate Lyase Preliminary experiments were carried out with immobilized CL.The resulting probe showed poor reproducibility and a marked loss of activity with repeated injections of citric acid, eliminating the advantages from the immobilization of the other enzymes. Planta et al.10 reported a 50% loss of the initial activity after 15–20 sample injections. Magnesium complexes of the enolic form of oxaloacetate appear to be responsible for the inactivation of the enzyme.15 The effect of the CL on the response was investigated by varying the enzyme additions (0–0.6 U ml21) to 0.15 mm citric acid standards.The saturation point is 0.4 U ml21. The same profile was also recorded when a standard solution of 0.4 mm citric acid was used. The Zn2+– enol oxaloacetate complex is less inhibitory15 and so the effect of ZnCl2 on the activity of CL was examined. Addition of ZnCl2 to standard solutions up to 0.03 mm showed no effect on the response. In contrast, at concentrations up to 0.13 mm a decrease in the response (approximately 8–10%) was observed as Zn2+ is also an inhibitor of OACD.25 At concentrations higher than 0.2 mm the formation of a Zn3(PO4)2 precipitate was evident. Interferences Interference by metal ions, amino acids and other organic acids present in real samples was investigated by applying the method of mixed solutions in the presence of 0.1 mm citric acid.Interferants were added at concentrations much higher than those in the real samples after dilution.The effect on the relative response is shown in Table 3, where only for malic acid was a small increase in the signal observed, presumably owing to its structural similarity with oxaloacetic acid. Because of the cellulose acetate membrane there is no interference effect from ascorbic acid. Application to Standards and Real Samples Under the optimum conditions, a series of calibration graphs, current/nA = f([analyte/ mm]), were constructed, applying the least-squares method. Using the Immunodyne ABC membrane Fig. 3 pH profile of the pyruvate (5) and oxaloacetate (-) biosensors, using the Immunodyne ABC membranes, 0.3 mm substrate; Parameters: 70 mm MOPS; 1 mm KH2PO4; 2 mm MgCl2; 1 mm TPP. Fig. 4 Flow rate profile of the pyruvate (5) and oxaloacetate (-) biosensors, using the Immunodyne ABC membranes; 0.3 mm substrate; Parameters: 70 mm MOPS, pH 7.5; 1 mm KH2PO4; 2 mm MgCl2; 1 mm TPP. 1104 Analyst, October 1997, Vol. 122(5 U POD per side), a linear relationship was obtained between the response and the pyruvate concentration in the range 0.01–0.9 mm with a correlation coefficient, r = 0.999. Data fitted the equation y = (20.02 ± 0.51) + (136.27 ± 1.27) [pyruvate]. The detection limit was 5 mm pyruvate for a signalto- noise ratio of 3 (S/N = 3). By using Immunodyne ABC and Biodyne B membranes, two calibration graphs, linear over the concentration range 0.015–0.6 mm oxaloacetate, were plotted.The equations for the straight lines were y = (0.18 ± 0.43) + (131.10 ± 1.53) [oxaloacetate], and y = (0.303 ± 1.07) + (106.77 ± 3.71) [oxaloacetate], with correlation coefficients r = 0.999 and r = 0.998, respectively. The detection limits (S/N = 3) were 4 and 10 mm oxaloacetate, respectively. By applying these graphs, pyruvate and oxaloacetate were determined in standard solutions and the mean relative error was 1.8% and 2.1%, respectively. Using the Immunodyne ABC membrane and citric acid standards, a calibration graph, linear over the range 0.015-0.5 mm, with a correlation coefficient r = 0.999, fitting the equation y = (0.09 ± 0.44) + (123.42 ± 2.67)[citrate], was constructed.The detection limit (S/N = 3) was 4 mm citrate and the RSD of the method was calculated as 1.0% (n = 8, 0.24 mm). Results are shown in Fig. 5. The proposed method was applied to fruits, juices and sport drinks for the determination of citric acid. The results for various samples are summarized in Table 4.Each sample required a minimum dilution of 1 + 99, whereas for orange and lemon juices a dilution of 1 + 449 and 1 + 1999, respectively, was required. The results were compared with those obtained with the Boehringer test kit. The mean relative error was 2.4%. The accuracy of the method was also verified by recovery studies performed by adding standard citric acid solutions to samples. According to the literature, apple and avocado do not contain citric acid26 and this was also verified with the proposed method.Recoveries of 96–105% were achieved, as shown in Table 5. The authors thank the EC (Project: MAT-1.ST93-0034) for financial support. Thanks are also extended to L. Arbizzani, from Pall Italia srl, who kindly donated samples of the membranes. M. I. P. thanks Professor G. Palleschi for valuable advice during his visit to ‘Tor Vergata’, University of Rome, and the European Science Foundation (Programme ABI).References 1 Gardner, W. H., in Handbook of Food Additives, ed. Furioc, T. E., CRC Press; Cleveland, OH, 2nd edn., 1972, pp. 242–246. 2 Murthy, G. K., Masurowsky, E. B., Campbell, J. E., and Edmondson, L. F., US Pat. 3 020 161, 1962; Chem. Abstr., 1962, 56, P14682g. 3 Kasai, Y., Tanimura, T., and Tamura, Z., Anal. Chem., 1975, 47, 34. 4 Coppola, E. D., Conrad, E. C., and Cotter, R., J. Assoc. Off. Anal. Chem., 1978, 61, 1490. 5 Bocek, P., Lekova, K., Deml, M., and Janak, J., J. Chromatogr., 1976, 117, 97. 6 Matsumoto, K., Ishida, K., Nomura, T., and Osajima, Y., Agric. Biol. Chem., 1984, 48, 2211. 7 Dunemann, L., Anal. Chim. Acta, 1989, 221, 19. 8 Spector, L. B., in The Enzymes, ed. Boyer. P. D., Academic Press, New York, 3rd edn., 1975, vol. VIII, 378. 9 Moellering, H., and Gruber, W., Anal. Biochem.., 1966, 17, 369. Table 3 Interference effect of various compounds on the assay of citric acid. The values in parentheses are the concentrations of the compounds in mm.All solutions contained 0.1 mm citric acid and were compared with the activity of plain 0.1 mm citric acid taken as 100% Relative Interferant activity (%) None 100 Potassium (5) 101 Sodium (5) 100 Alanine (5) 98.5 Lysine (5) 98.5 Leucine (5) 99 Glutamic acid (5) 99.5 Lactic acid (2) 101.5 Adipic acid (2) 99.5 Tartaric acid (2) 103.5 Butyric acid (2) 101.5 Malic acid (2) 106 Ascorbic acid (1) 101 Acetic acid (2) 100 Oxalic acid (2) 100 Isocitric acid (2) 101 Fig. 5 Calibration graph of citric acid with all the parameters optimized (see text). FI traces top left, reproducibility of the system (0.24 mm citric acid, n = 8). Bottom right, calibration graph for citric acid. Peaks 2–10 correspond to concentrations within the linear range while peak 1 represents a concentration of 0.007 mm citric acid. Table 4 Determination of citric acid in various real samples. The standard deviation of the mean ranges from 0.01 to 0.09 mm Proposed Reference Relative Dilution method*/ method†/ error Sample ratio mm mm (%) Lemonade (IVI) 10 2.24 2.26 20.9 Ice-tea lemon 10 1.11 1.16 24.3 Lemon juice 100 2.41 2.38 +1.3 Juice (Florina) 10 3.50 3.65 24.1 Lucozade sport 10 3.00 3.05 21.7 Orange juice 50 1.66 1.63 +1.8 * Average of three runs.† Boehringer–Mannheim test kit. Table 5 Recovery of citric acid added to real samples Sample Added/1024 m Found/1024 m Recovery (%) Lemonade (IVI) 0.60 0.58 96 Juice (Florina) 0.80 0.84 105 Lucozade sport 0.70 0.68 98 Ice-tea lemon 0.80 0.78 98 Lemon juice 0.60 0.62 103 Apple (5%) 0 0 — Apple (5%) 17.11 17.70 104 Avocado (5%) 0 0 — Avocado (5%) 17.11 16.61 97 Analyst, October 1997, Vol. 122 110510 Plant�a, M., Lazaro, F., Puchades, R., and Maquieira, A., Analyst, 1993, 118, 1193. 11 Hasebe, K., Hikima, S., Kakizaki, T., and Yoshida, H., Fresenius’ Z. Anal. Chem., 1989, 333, 19. 12 Hikima, S., Hasebe, K., and Taga, M., Electroanalysis, 1992, 4, 801. 13 Matsumoto, K., Tsukatani, T., and Okajima, Y., Electroanalysis, 1995, 7, 527. 14 Gajovic, N., Warsinke, A., and Scheller, F. W., J. Chem. Tech. Biotechnol., 1995, 63, 337. 15 Dagley, S., in Methods in Enzymology, ed. Lowestein, I. M., Academic Press, New York, 1969, vol. XIII, ch. 67. 16 Prodromidis, M. I., Tsibiris, A. B., and Karayannis, M. I., J. Autom. Chem., 1995, 17, 187. 17 Enzymatic Analysis. A Practical Guide, ed. Passonneau, J. V., and Lowry, H. O., Humana Press, Clifton, NJ, 1993. 18 Assoland-Vinet, C.H., and Coulet, P. R., Anal. Lett., 1986 , 19, 875. 19 Mizutani, F., Tsuda, K., Karube, I., Suzuki, S., and Matsumoto, K., Anal. Chim. Acta, 1980, 118, 65. 20 Mascini, M., Iannello, M., and Palleschi, G., Anal. Chim. Acta, 1983, 146, 135. 21 Palleschi, G., Nabi Rahni, M. A., Lubrano, G. J., Ngwainbi, J. H., and Guilbault, G. G., Anal. Biochem., 1986, 159, 114. 22 Albery, W. J., Bartlett, P. N., and Craston, D. H., J. Electroanal. Chem., 1985, 194, 223. 23 Kihara, K., Yasukawa, E., and Hirose, S., Anal.Chem., 1984, 56, 1876. 24 Hager, L. P., and Lipmann F., in Methods in Enzymology, ed. Colowick, S. P., and Kaplan, N. O., Academic Press, New York, 1955, vol. I, ch. 75. 25 Jetten, M. S. M., and Sinskey, A. J., Antonie van Leeuwenhoek Int. J., 1995, 67, 221; Chem. Abstr., 123; 283169. 26 Joslyn, M. A., in Methods In Food Analysis, Academic Press; New York, 1970, ch. XIV, p. 408. Paper 7/02312J Received April 4, 1997 Accepted June 23, 1997 1106 Analyst, October 1997, Vol. 122 Bioelectrochemical Determination of Citric Acid in Real Samples Using a Fully Automated Flow Injection Manifold Mamas I.Prodromidisa, Stella M. Tzouwara-Karayannia, Miltiades I. Karayannis*a and Pankaj M. Vadgamab a University of Ioannina, Laboratory of Analytical Chemistry, Ioannina 45110, Greece b University of Manchester, Department of Medicine, Hope Hospital, Salford, UK M6 HD An enzymic method for the determination of citric acid in fruits, juices and sport drinks is proposed.The method is based on the action of the enzymes citrate lyase, oxaloacetate decarboxylase and pyruvate oxidase, which convert citric acid into H2O2 with the latter being monitored amperometrically with a H2O2 probe. The enzymes pyruvate oxidase and oxaloacetate decarboxylase were immobilized. A multi-membrane system, consisting of a cellulose acetate membrane for the elimination of interferants, an enzymic membrane and a protective polycarbonate membrane were placed on a Pt electrode and used with a fully automated flow injection manifold.Several parameters were optimized, resulting in a readily constructed and reproducible biosensor. Interference from various compounds present in real samples was minimized. Calibration graphs were linear over the range 0.01–0.9 mm pyruvate, 0.015–0.6 mm oxaloacetate and 0.015–0.5 mm citrate. The throughput was 30 samples h21 with an RSD of 1.0% (n = 8); the mean relative error was 2.4% compared with a standard method.The recovery was 96–104%. A 8–10% loss of the initial activity of the sensor was observed after 100–120 injections. Keywords: Amperometric citrate determination; multimembrane biosensor; flow injection analysis; fruits, juices and sport drinks Citric acid is present in numerous natural products and is a key tribasic acid involved in both plant and animal aerobic respiration. Several fresh fruits such as lemons and limes owe their sharp taste to the presence of the citrate anion.1 Citric acid is also an additive in industry, mainly as a preservative and an acidulant.It is added to some dairy products to improve and protect both flavor and aroma. It has been used for radioactive Sr2+ from suspect milk after radiation fallout, and has been useful for chelating trace metals which can cause haze or deterioration of color and flavor.2 Several methods have been proposed for the determination of citric acid, based on ion-exchange chromatography,3 HPLC4 and isotachophoresis;5 these are time consuming procedures, and sample clean-up is required in order to separate citric acid from other co-existing tricarboxylic acids.Other approaches based on conductimetric6 or spectrophotometric methods7 suffer from selectivity as they are based on non-specific reactions with carboxylic acids. The need for simple separation could be potentially solved, by using a high specificity enzyme such as citrate lyase8 (CL). Moellering and Gruber9 reported a method with soluble CL, lactate dehydrogenase and malate dehydrogenase (MDH), and monitored the decrease in the absorbance of the NADH at 340 nm.Although relatively slow and expensive in enzyme the approach works well, as a batch method, and is widely used in routine off-line analysis. Since then several other approaches have been proposed; some make use of soluble CL and immobilized MDH with NADH10 detection, some use CL and oxaloacetate decarboxylase (OACD) in soluble11 or immobilized12 form in conjunction with polarography.The direct amperometric determination of citric acid has been proposed13 in connection with an ascorbate oxidase reactor to eliminate ascorbic acid interference. Gajovic et al.14 used CL, purified by recrystallisation and ultrafiltration and then entrapped in gelatin. This method needs recalibration every six runs probably owing to the gradual loss of the activity of the CL.The enzyme could be reactivated after 1 h incubation with adenosine 5A- triphosphate (ATP) and CH3COOH. Measurements using a non-enzymic membranwere a proposed route to the elimination of interferent species. CL is usually used in soluble form because of ready inactivation by complex formation with Mg2+ and Zn2+ and also the enol-form of oxaloacetate,15 itself the product of its action on citrate. The proposed method is based on a sequence of reactions involving thiamine pyrophosphate (TPP) and the enzymes CL, OACD and pyruvate oxidase (POD), according to the following scheme: citrate oxaloacetate CH COOH CL 3 ¾ ®¾ + (1) oxaloacetate pyruvate + CO OACD 2 ¾ ® ¾¾ (2) pyruvate + HPO O acetyl-phosphate CO H O 4 2 2 2 2 POD TPP, Mg 2+ 2-+ ® + + (3) The final product H2O2 was monitored amperometrically by means of a Pt electrode, mounted on a wall-jet flow-through cell, polarised at +0.65 V versus an Ag–AgCl reference electrode.The multi-membrane biosensor gave an interferantfree system, with good analytical characteristics in terms of accuracy, reproducibility and operational simplicity, which is a key advantage of biosensors over conventional enzymic assays. Additionally, the system was combined with a flow injection (FI) manifold, fully automated by means of resident reported software.16 Experimental Apparatus This work was carried out using an in-house fully automated FI manifold.Electrochemical experiments were run using a computer-controlled potentiostat (Eco Chemie/Autolab, Utrecht, The Netherlands). Solutions were pumped through the manifold using a four-channel peristaltic pump (Gilson, Villiers Analyst, October 1997, Vol. 122 (1101–1106) 1101le Bel, France) and sample injections made with a pneumatically actuated injection valve (Rheodyne, Cotati, CA, USA). A resident program, synchronized with the acquisition software, ensured full control of pump and valves, eliminating manual manipulation; a detailed description of the program is given elsewhere.16 A three-electrode flow-through detector (Metrohm 656, Herisau, Switzerland) was used for the amperometric monitoring.This consists of a wall-jet thermostated cell (volume < 1 ml) with a Pt (diameter 1.6 mm, Bioanalytical Systems, W. Lafayette, IN, USA) working electrode, a built-in Au auxiliary electrode and an Ag–AgCl reference electrode. A schematic diagram of the system is shown in Fig. 1(A). Chemicals POD (EC 1.2.3.3, from Pediococcus sp., 43 U mg21), OACD (EC 4.1.1.3, from Pseudommonas sp., 265 U mg21), CL (EC 4.1.3.6, from Enterobacter aerogenes, 0.26 U mg21) all in lyophilized form and cellulose acetate (approximately 40% acetyl), were obtained from Sigma (St. Louis, MO, USA). 3- Morpholinopropanesulfonic acid (MOPS) and poly(vinyl acetate) (PVA, Mr 167 000 Da) were supplied by Aldrich (Gollingham, Germany). The enzymic test kit for citric acid determinations was purchased from Boehringer (Mannheim, Germany).All other (analytical-reagent grade) chemicals were purchased from Sigma. Solutions Working (MOPS 70 mm, pH 7.6) and immobilization (MOPS 50 mm, pH 7.35) buffer solutions were filtered through a 1.0 mm pore diameter microporous polycarbonate (PC) membrane (Millipore, Bedford, MA, USA), prior to use. Stock solutions of pyruvic (50 mm) and oxaloacetic acid (50 mm) were prepared by dissolving 27.5 and 33.1 mg of sodium pyruvate and oxaloacetic acid in 5.0 ml of 0.1 and 0.5 m HCl, respectively; this prevented their polymerization or decarboxylation.Both solutions were stored at 4 °C and prepared weekly, owing to their instability.17 A stock solution of TPP (20 mm, pH 6.2) was prepared by dissolving 181.4 mg of TPP in 20 ml of the working buffer. Enzyme solutions were prepared in the immobilization buffer, otherwise working buffer solution was used throughout. Membranes Polymeric nylon 66 membranes, thickness 120 mm, viz., Biodyne A (amphoteric, 50% amino, 50% carboxyl, 0.2 mm porosity), Biodyne B (positively charged with pore surfaces populated by a high density of quaternary ammonium groups, 0.45 mm porosity), Biodyne C (negatively charged, contains 100% carboxyl, 0.45 mm porosity) and Immunodyne ABC (preactivated, 0.2 mm porosity) were a kind gift from Pall Filtration (Milan, Italy).HA (mixed ester cellulose, 0.45 mm porosity) membranes were purchased from Millipore.PC membranes (thickness 10 mm, 0.03 and 0.05 mm porosity) and dialysis tubing (12 000 Mr cut-off) were supplied by Nuclepore (Cambridge, MA, USA) and Sigma, respectively. For casting cellulose acetate membranes a wet film applicator (5 in, 1–8 mils, URAI, Milan, Italy) was used. Preparation of the Enzymic Membranes POD was immobilized by ionic and/or covalent bonding onto nylon 66 membranes, following the spot wetting method:18 A 10 ml (10 U) aliquot of the POD solution was symmetrically pipetted onto each side of the dry membrane (diameter 6 mm) and left to air-dry for 5 min.Alternatively, physical entrapment of POD was carried out by passing 50 ml (10 U) of POD solution through an HA membrane (diameter 6 mm) while slight suction was applied.19 In both cases, unattached protein was removed by washing the membranes (3 3 10 min) with the immobilization buffer. POD and OACD were co-immobilized asymmetrically onto Biodyne B and Immunodyne ABC membranes, according to a procedure reported in the literature:20 A 10 ml (10 U) aliquot of the POD solution was applied onto the rough side of the dry membrane (1 3 1 cm2) and allowed to react for 2–3 min.Then, 6 ml (15 U) of the OACD solution were applied onto the smooth side of the membrane and allowed again to react for 15 min. The membranes were washed and stored in the immobilization buffer at +4 °C. Assembly of the Sensor The cellulose acetate membrane (20 mm thick, 100 Da cut-off) was first placed on the Pt surface to eliminate interference from electroactive species.21 This membrane was prepared in our laboratory by dissolving 3.996 g of cellulose acetate and 4 mg of PVA in a mixture of 60 ml of acetone and 40 ml of cyclohexanone (after a minor modification of the procedure reported by Palleschi et al.21).Then the membrane bearing the enzyme(s) was superimposed with an outer PC membrane in order to prevent microbial attack and also leaching of enzyme.With the oxaloacetate biosensor, the rough surface of the membrane was directed towards the cellulose acetate; POD membranes could be placed with any orientation. All the Fig. 1 (A) Schematic representation of FI manifold employed for citric acid determinations: C, carrier; R, TPP reagent. (B) Assembly of the biosensor; LM, large molecules; AN, analyte; IN, interferants; Pt, platinum electrode. CA, cellulose acetate; EM, enzymic membrane; PC, polycarbonate membrane.(C) i, POD membrane; ii, random application; and iii, asymmetric application. 1102 Analyst, October 1997, Vol. 122membranes were tightly fitted over the electrode with the aid of an O-ring [Fig. 1(B)]. PC membranes of different porosity (0.03 and 0.05 mm) and a dialysis membrane (modified with 1% Na2CO3 and 10 mm Trizma–1 mm EDTA22) were alternatively used as protective membranes. After preliminary evaluation of the effect of these membranes on the pyruvate biosensor, in terms of response times, sensitivity, linearity and operational stability, a porosity of 0.03 mm was finally selected.Preparation of Sample Solutions A 5 g amount of fruit sample was homogenized with a blender, diluted in 100 ml of the working buffer and finally filtered through a 1.0 mm PC membrane. Juice and sport drink samples were directly diluted in 10 ml of working buffer, while carbonated samples were sonicated (1 min, 100 W) prior to dilution.Procedure The carrier (70 mm MOPS of pH 7.6 containing 3.22 nm MgCl2 and 1.61 mm KH2PO4) and TPP [2.64 mm, pH 6.2 (2.64 ml of 20 mm TPP in 20 ml of water)] streams were continuously pumped at flow rates of 0.23 and 0.14 ml min21, respectively, towards the probe until a stable baseline current was reached (1–2 nA, within 10–15 min). Standard or sample solutions of citric acid [40 ml (0.4 U) CL + x ml sample + (960 2x) ml carrier] were introduced as short pulses of 120 ml via the loop injection valve. For the oxaloacetate or pyruvate biosensor, the composition of the standards was [x ml standard + (5000 2 x) ml carrier].The peak height of the current response was taken as a measure of the analyte concentration. Results and Discussion Enzyme Membranes Commencing from the first enzymic step [eqn. (3)], POD was immobilized onto several commercially available membranes. The wide range of the chemical and physical properties of the membranes led us to find the most efficient matrix for the construction of the pyruvate biosensor.As well as the membrane spot wetting immobilization, membrane immersion18 was tested using a 0.32 mg per 2 ml (10 U POD) enzymic solution; the latter gave poor immobilization efficiencies. Enzyme loading was tested separately for POD and OACD and defined the enzyme loading necessary to obtain diffusional limitation of response, i.e., the response maximum.14 Saturation study of the membrane was made using the Immunodyne ABC membrane; this is claimed to have the greatest binding capacity.The response to pyruvate with different enzyme loading solutions (2–12 U POD) was studied (Fig. 2). Experiments were then performed, applying the useful enzyme content (10 U POD) onto different membranes. The relative apparent measurable efficiencies, which reflect the biocatalytic efficiency of the immobilized enzymes, are shown in Table 1. Biodyne B and Immunodyne ABC membranes showed the best results and so further work was carried out using these membranes. By keeping the amount of POD at the optimum value, further membranes were prepared by varying the amount of the applied OACD from 5 to 20 U.The responses of oxaloacetate biosensors with different enzyme loadings are shown in Fig. 2. Asymmetric co-immobilization of POD and OACD gave greater current signals than those with random application of the same amount of the enzymes [Fig. 1(C)]. Probe Lifetime POD is a relatively unstable enzyme probably owing to the gradual loss of its cofactor flavin adenine dinucleotide (FAD), during both the immobilization procedure and the measurements.The addition of several compounds such as FAD, TPP, MgCl2, glycerol and (NH4)2SO4 to the immobilization or storage buffer did not improve either the immobilization efficiency or the long-term stability. Indeed, this led to a more complicated storage of the membranes and greater cost. OACD is a more stable enzyme; therefore, lifetime experiments were carried out with the pyruvate biosensor (Table 2).The proposed approach is more economical than other attempts, with similar13 or slightly better lifetimes23 where the preparation time of the enzymic membranes and the amount of the enzymes are significantly greater ( > 30 h preparation time and 10–26-fold amounts of the enzyme). Concentration of Activators and Cofactors It has been reported24 that the activity of POD is affected by the presence of inorganic phosphate, divalent cations such as Mg2+, Ca2+, Mn2+ or Co2+ and cofactors such as TPP and FAD.This optimization was carried out with the probe assembled as a pyruvate biosensor, using 0.5 mm pyruvate solution as a sample. Fig. 2 Enzyme loading test using the Immunodyne ABC membrane: 5 POD (2–12 U); -, POD (10 U) + OACD (5–20 U); 0.5 mm substrate. Parameters: 70 mm MOPS; pH 7.5; 1 mm MgCl2; 2 mm TPP; 1 mm KH2PO4. Table 1 Relative efficiencies of various polymeric membranes used as supports for the immobilization of POD. 0.3 mm pyruvate; 10 U POD Membrane Current/nA RE (%)* Biodyne A 29.3 68 Biodyne B 42.8 100 Biodyne C 27.3 64 Immunodyne ABC 40.7 95 Millipore HA 22.1 52 * Relative efficiency with respect to the most active membrane. Table 2 Lifetime of the pyruvate biosensor. Immunodyne ABC membrane; 10 U POD; 0.2 mm pyruvate* Day Current/nA Activity (%) 1 26.0 100 2 23.4 90 3 20.7 80 4 19.0 73 5 9.2 35 6 5.5 21 7 1.0 4 * Continuous use: 100–200 injections (8–10% loss of the initial activity). Use after storage: minimum 15 injections per day.Analyst, October 1997, Vol. 122 1103TPP was initially included in the carrier stream, but it was found to be unstable in this environment (decay of signal output), and so was pumped separately [Fig. 1(A)] achieving, furthermore, better mixing with the sample solution. In the absence of TPP no response was observed while for concentrations higher than 0.8 mm TPP the response became constant; subsequent work used 1mM TPP.Several cations were examined as activators of POD. In addition to sensitivity, the criterion for the final selection was the stability of the sensor. Since Ca2+ and Co2+ precipitate with phosphate they must be avoided; Ca2+ also competed with the binding site for Mg2+, vital for CL activity. A 2 mm MgCl2 concentration was found to be optimum for the performance of the biosensor. Inorganic phosphate, as KH2PO4, was tested in the concentration range 0–2 mm and a concentration of 1 mm was selected; the graph was similar to that seen with Mg2+.FAD in the carrier stream (up to 0.5 mm) had no effect on the current output; it is likely that the enzyme preparation contains sufficient FAD for the catalytic reaction and no further FAD was needed. Using POD–OACD membranes and 0.15 mm oxaloacetate, the effect of TPP, MgCl2 and KH2PO4 was re-examined and similar optima were observed.The effect of Mn was further investigated and it was found to increase the activity of the system (approximately 6–8%) when a concentration of 0.1 mm MnCl2 was added to the carrier stream. At higher concentrations a progressive decrease of the signal was observed, accompanied by the formation of Mn3(PO4)2; hence, subsequent work was carried out in the absence of MnCl2. Working Conditions The pH of the working buffer was also investigated in several buffering systems, such as MOPS, glycylglycine and Trizma- HCl, covering the pH range 7–8.The last gave lower responses, probably owing to its reaction with Mg2+ and oxaloacetate ions. Using POD–OACD membranes, sequential injections of 0.3 mm pyruvate and 0.3 mm oxaloacetate, at different pH values of 70 mm MOPS, were performed. The optimum pH for both the pyruvate and oxaloacetate biosensors is 7.5 as shown in Fig. 3. The numbers that appear on the graph were calculated as (S1/S2) 3 100 (where S1 and S2 represent the signals obtained for standards of 0.3 mm oxaloacetate and pyruvate, respectively) and reflect the efficiency of the conversion of oxaloacetate to pyruvate.The efficiencies of the conversion were also investigated at different flow rates (reaction times), following the procedure given above. Flow rate profiles are shown in Fig. 4. An overall flow rate of 0.37 ml min21 was finally selected, which reconciles fairly high peaks and satisfactory sample throughput (30 h21).A sample volume of 120 ml was used as it prevented peak broadening (dispersion coefficient 1.22–1.25) and also ensured high sensitivity. The sensitivity of the oxaloacetate biosensor also increased with the temperature, levelling off at a maximum value of 47 °C. Above this temperature thermal inactivation dominates over the increase in the collision frequency, resulting in a decrease of the signal. All experiments were carried out at 30 °C, where the stability of the biosensor was the same as at room temperature.Amount of Citrate Lyase Preliminary experiments were carried out with immobilized CL. The resulting probe showed poor reproducibility and a marked loss of activity with repeated injections of citric acid, eliminating the advantages from the immobilization of the other enzymes. Planta et al.10 reported a 50% loss of the initial activity after 15–20 sample injections. Magnesium complexes of the enolic form of oxaloacetate appear to be responsible for the inactivation of the enzyme.15 The effect of the CL on the response was investigated by varying the enzyme additions (0–0.6 U ml21) to 0.15 mm citric acid standards.The saturation point is 0.4 U ml21. The same profile was also recorded when a standard solution of 0.4 mm citric acid was used. The Zn2+– enol oxaloacetate complex is less inhibitory15 and so the effect of ZnCl2 on the activity of CL was examined. Addition of ZnCl2 to standard solutions up to 0.03 mm showed no effect on the response.In contrast, at concentrations up to 0.13 mm a decrease in the response (approximately 8–10%) was observed as Zn2+ is also an inhibitor of OACD.25 At concentrations higher than 0.2 mm the formation of a Zn3(PO4)2 precipitate was evident. Interferences Interference by metal ions, amino acids and other organic acids present in real samples was investigated by applying the method of mixed solutions in the presence of 0.1 mm citric acid.Interferants were added at concentrations much higher than those in the real samples after dilution. The effect on the relative response is shown in Table 3, where only for malic acid was a small increase in the signal observed, presumably owing to its structural similarity with oxaloacetic acid. Because of the cellulose acetate membrane there is no interference effect from ascorbic acid. Application to Standards and Real Samples Under the optimum conditions, a series of calibration graphs, current/nA = f([analyte/ mm]), were constructed, applying the least-squares method.Using the Immunodyne ABC membrane Fig. 3 pH profile of the pyruvate (5) and oxaloacetate (-) biosensors, using the Immunodyne ABC membranes, 0.3 mm substrate; Parameters: 70 mm MOPS; 1 mm KH2PO4; 2 mm MgCl2; 1 mm TPP. Fig. 4 Flow rate profile of the pyruvate (5) and oxaloacetate (-) biosensors, using the Immunodyne ABC membranes; 0.3 mm substrate; Parameters: 70 mm MOPS, pH 7.5; 1 mm KH2PO4; 2 mm MgCl2; 1 mm TPP. 1104 Analyst, October 1997, Vol. 122(5 U POD per side), a linear relationship was obtained between the response and the pyruvate concentration in the range 0.01–0.9 mm with a correlation coefficient, r = 0.999. Data fitted the equation y = (20.02 ± 0.51) + (136.27 ± 1.27) [pyruvate]. The detection limit was 5 mm pyruvate for a signalto- noise ratio of 3 (S/N = 3). By using Immunodyne ABC and Biodyne B membranes, two calibration graphs, linear over the concentration range 0.015–0.6 mm oxaloacetate, were plotted.The equations for the straight lines were y = (0.18 ± 0.43) + (131.10 ± 1.53) [oxaloacetate], and y = (0.303 ± 1.07) + (106.77 ± 3.71) [oxaloacetate], with correlation coefficients r = 0.999 and r = 0.998, respectively. The detection limits (S/N = 3) were 4 and 10 mm oxaloacetate, respectively. By applying these graphs, pyruvate and oxaloacetate were determined in standard solutions and the mean relative error was 1.8% and 2.1%, respectively. Using the Immunodyne ABC membrane and citric acid standards, a calibration graph, linear over the range 0.015-0.5 mm, with a correlation coefficient r = 0.999, fitting the equation y = (0.09 ± 0.44) + (123.42 ± 2.67)[citrate], was constructed. The detection limit (S/N = 3) was 4 mm citrate and the RSD of the method was calculated as 1.0% (n = 8, 0.24 mm).Results are shown in Fig. 5. The proposed method was applied to fruits, juices and sport drinks for the determination of citric acid.The results for various samples are summarized in Table 4. Each sample required a minimum dilution of 1 + 99, whereas for orange and lemon juices a dilution of 1 + 449 and 1 + 1999, respectively, was required. The results were compared with those obtained with the Boehringer test kit. The mean relative error was 2.4%. The accuracy of the method was also verified by recovery studies performed by adding standard citric acid solutions to samples. According to the literature, apple and avocado do not contain citric acid26 and this was also verified with the proposed method.Recoveries of 96–105% were achieved, as shown in Table 5. The authors thank the EC (Project: MAT-1.ST93-0034) for financial support. Thanks are also extended to L. Arbizzani, from Pall Italia srl, who kindly donated samples of the membranes. M. I. P. thanks Professor G. Palleschi for valuable advice during his visit to ‘Tor Vergata’, University of Rome, and the European Science Foundation (Programme ABI).References 1 Gardner, W. H., in Handbook of Food Additives, ed. Furioc, T. E., CRC Press; Cleveland, OH, 2nd edn., 1972, pp. 242–246. 2 Murthy, G. K., Masurowsky, E. B., Campbell, J. E., and Edmondson, L. F., US Pat. 3 020 161, 1962; Chem. Abstr., 1962, 56, P14682g. 3 Kasai, Y., Tanimura, T., and Tamura, Z., Anal. Chem., 1975, 47, 34. 4 Coppola, E. D., Conrad, E. C., and Cotter, R., J. Assoc. Off.Anal. Chem., 1978, 61, 1490. 5 Bocek, P., Lekova, K., Deml, M., and Janak, J., J. Chromatogr., 1976, 117, 97. 6 Matsumoto, K., Ishida, K., Nomura, T., and Osajima, Y., Agric. Biol. Chem., 1984, 48, 2211. 7 Dunemann, L., Anal. Chim. Acta, 1989, 221, 19. 8 Spector, L. B., in The Enzymes, ed. Boyer. P. D., Academic Press, New York, 3rd edn., 1975, vol. VIII, 378. 9 Moellering, H., and Gruber, W., Anal. Biochem.., 1966, 17, 369. Table 3 Interference effect of various compounds on the assay of citric acid.The values in parentheses are the concentrations of the compounds in mm. All solutions contained 0.1 mm citric acid and were compared with the activity of plain 0.1 mm citric acid taken as 100% Relative Interferant activity (%) None 100 Potassium (5) 101 Sodium (5) 100 Alanine (5) 98.5 Lysine (5) 98.5 Leucine (5) 99 Glutamic acid (5) 99.5 Lactic acid (2) 101.5 Adipic acid (2) 99.5 Tartaric acid (2) 103.5 Butyric acid (2) 101.5 Malic acid (2) 106 Ascorbic acid (1) 101 Acetic acid (2) 100 Oxalic acid (2) 100 Isocitric acid (2) 101 Fig. 5 Calibration graph of citric acid with all the parameters optimized (see text). FI traces top left, reproducibility of the system (0.24 mm citric acid, n = 8). Bottom right, calibration graph for citric acid. Peaks 2–10 correspond to concentrations within the linear range while peak 1 represents a concentration of 0.007 mm citric acid. Table 4 Determination of citric acid in various real samples. The standard deviation of the mean ranges from 0.01 to 0.09 mm Proposed Reference Relative Dilution method*/ method†/ error Sample ratio mm mm (%) Lemonade (IVI) 10 2.24 2.26 20.9 Ice-tea lemon 10 1.11 1.16 24.3 Lemon juice 100 2.41 2.38 +1.3 Juice (Florina) 10 3.50 3.65 24.1 Lucozade sport 10 3.00 3.05 21.7 Orange juice 50 1.66 1.63 +1.8 * Average of three runs. † Boehringer–Mannheim test kit. Table 5 Recovery of citric acid added to real samples Sample Added/1024 m Found/1024 m Recovery (%) Lemonade (IVI) 0.60 0.58 96 Juice (Florina) 0.80 0.84 105 Lucozade sport 0.70 0.68 98 Ice-tea lemon 0.80 0.78 98 Lemon juice 0.60 0.62 103 Apple (5%) 0 0 — Apple (5%) 17.11 17.70 104 Avocado (5%) 0 0 — Avocado (5%) 17.11 16.61 97 Analyst, October 1997, Vol. 122 110510 Plant�a, M., Lazaro, F., Puchades, R., and Maquieira, A., Analyst, 1993, 118, 1193. 11 Hasebe, K., Hikima, S., Kakizaki, T., and Yoshida, H., Fresenius’ Z. Anal. Chem., 1989, 333, 19. 12 Hikima, S., Hasebe, K., and Taga, M., Electroanalysis, 1992, 4, 801. 13 Matsumoto, K., Tsukatani, T., and Okajima, Y., Electroanalysis, 1995, 7, 527. 14 Gajovic, N., Warsinke, A., and Scheller, F. W., J. Chem. Tech. Biotechnol., 1995, 63, 337. 15 Dagley, S., in Methods in Enzymology, ed. Lowestein, I. M., Academic Press, New York, 1969, vol. XIII, ch. 67. 16 Prodromidis, M. I., Tsibiris, A. B., and Karayannis, M. I., J. Autom. Chem., 1995, 17, 187. 17 Enzymatic Analysis. A Practical Guide, ed. Passonneau, J. V., and Lowry, H. O., Humana Press, Clifton, NJ, 1993. 18 Assoland-Vinet, C. H., and Coulet, P. R., Anal. Lett., 1986 , 19, 875. 19 Mizutani, F., Tsuda, K., Karube, I., Suzuki, S., and Matsumoto, K., Anal. Chim. Acta, 1980, 118, 65. 20 Mascini, M., Iannello, M., and Palleschi, G., Anal. Chim. Acta, 1983, 146, 135. 21 Palleschi, G., Nabi Rahni, M. A., Lubrano, G. J., Ngwainbi, J. H., and Guilbault, G. G., Anal. Biochem., 1986, 159, 114. 22 Albery, W. J., Bartlett, P. N., and Craston, D. H., J. Electroanal. Chem., 1985, 194, 223. 23 Kihara, K., Yasukawa, E., and Hirose, S., Anal. Chem., 1984, 56, 1876. 24 Hager, L. P., and Lipmann F., in Methods in Enzymology, ed. Colowick, S. P., and Kaplan, N. O., Academic Press, New York, 1955, vol. I, ch. 75. 25 Jetten, M. S. M., and Sinskey, A. J., Antonie van Leeuwenhoek Int. J., 1995, 67, 221; Chem. Abstr., 123;83169. 26 Joslyn, M. A., in Methods In Food Analysis, Academic Press; New York, 1970, ch. XIV, p. 408. Paper 7/02312J Received April 4, 1997 Accepted June 23, 1997 1106 Analyst, October 1997, Vol. 122

ISSN:0003-2654    

DOI:10.1039/a702312j

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Detection of 2,4-Dichlorophenoxyacetic Acid Using a Fluorescence Immunoanalyzer Kim R. Rogers*a, Steven D. Kohla, Lee A. Riddicka and Thomas Glassb a US Environmental Protection Agency, National Exposure Research Laboratory, Las Vegas, NV 89193, USA b Sapidyne Inc., Boise, ID 83706, USA A flow immunoassay method for the measurement of 2,4-dichlorophenoxyacetic acid (2,4-D) was developed. The competitive fluorescence immunoassay relies on the use of antibody- or antigen-coated poly(methyl methacrylate) particles (98 mm diameter) as a renewable solid phase.The assay exhibits a dynamic range of 0.1–100 mg l21 using a monoclonal antibody or alternatively 10 mg l21 to 10 mg l21 using commercially available antiserum. The assay is demonstrated in buffered saline solution as well as in aquatic environmental media. The relative errors for the environmental matrices were similar to those for the buffer control. The precision of concentration values calculated at 1 mg l21 (for the assay using antiserum) were ±0.28, ±0.27 and ±0.43 mg l21 for the buffer, well water and river water matrices, respectively.The method shows cross-reactivity with compounds of closely related structure but little cross-reactivity with compounds dissimilar in structure to 2,4-D. The proposed automated competitive immunoassay method is rapid (between 7 and 15 min per assay), simple and potentially portable. Keywords: 2,4-Dichlorophenoxyacetic acid; fluoroimmunoassay; KinExA immunoanalyzer Methods for the rapid and cost-effective detection of pesticides in environmental settings have gained considerable attention in recent years, primarily due to concerns related to potential human exposure from spray run-off and spills into groundwater sources.1 Classical laboratory-based methods used to measure these compounds are typically time consuming and expensive.These methods usually require extensive extraction, derivatization and separation by GC or HPLC.To meet the need for faster and more cost-effective methods, a variety of techniques have recently been reported and a number of commercial products (methods) have been introduced. These methods are, for the most part, immunochemically based enzyme-linked immunosorbent assays (ELISA) using absorbance, fluorescence or electrochemical detection. These assays have typically been formatted for microtiter plates,2 thick film-based planar microwells, 3 microflow cells,4 test kits,5 or test tube-type optical spectrometers.6 Although automation has been incorporated into a number of these method formats, they typically require manual addition of reagents, several incubation steps, and physical manipulations such as changing tubes, trays or capillaries.Portable automated systems that avoid these physical manipulations and generate (as waste) only spent reagents show potential cost, time and waste disposal advantages for certain applications.Several approaches have been used for the automated immunoassay flow systems. Formats for these systems involve either regeneration of the antibody immobilized to the solid phase (i.e., disruption of antibody– antigen binding)7 or renewal of a mobile solid support (e.g., microspheres which are trapped in a flow cell) which has been coated with either the antibodies or antigens,8,9 or enzymes.10 The use of a mobile solid phase which is renewable for each assay offers certain technical advantages resulting in a fast, versatile and inexpensive assay system. 2,4-Dichlorophenoxyacetic acid (2,4-D) is widely used as a systemic herbicide for broad leaf weeds in a variety of crop and non-crop applications. This pesticide is typically monitored in environmental samples using GC methods which require extensive clean-up and derivatization, however, a number of potential field analytical methods have been reported. These methods include: ELISA3,4,9,11 enzyme immunoassay,1 immunoagglutination, 12 fluorescence polarization,13 and enzyme assays14 as well as several antibody-based biosensor methods. 15–17 Although most of these methods are portable and relatively simple to execute, they typically require multiple steps and generate solid wastes (e.g., plates, tubes and vials). In this paper, we describe the use of the KinExA immunoanalyzer to develop competitive fluorescence immunoassay methods for the detection of 2,4-D using both monoclonal and polyclonal antibodies.These methods use a mobile–renewable solid phase (i.e., microspheres) and can be fully automated. Experimental Instrumentation and KinExA Method The KinExA instrument is an automated fluorescence immunoassay system that uses microspheres as the solid phase (Sapidyne, Boise, ID, USA). For the assay formats described here, these beads [poly(methyl methacrylate), PMMA] were coated with either antibody or antigen and then packed into a capillary flow cell that is integrated into an epi-illumination filter fluorimeter system.For the format using antibody-coated beads, fluorescently labeled probe is introduced into the flow cell in the presence or absence of analyte and continuous interrogation of the bead column for analyte-sensitive accumulation of fluorescently labeled tracer on the solid phase forms the basis of this assay. For the format using antigen-coated beads, monoclonal antibody or anti-2,4-D antiserum is incubated with the analyte (2,4-D), then introduced into the flow cell containing antigencoated beads.This step is followed by fluorescently labeled secondary (species-specific, anti-IgG) antibody. For both formats, the observed fluorescence signal varies inversely with the analyte concentration. The competitive immunoassay format using the antibodycoated beads is described in Fig. 1(a). In this assay, the PMMA particles were coated with anti-sheep IgG which was used to capture and orient the primary antibody from sheep anti-2,4-D serum. Next, inhibition in the accumulation of fluoresceinlabeled 2,4-D tracer resulting from the presence of various concentrations of the analyte 2,4-D allowed the detection and quantification of this analyte.Another competitive immunoassay format described in Fig. 1(b) used beads coated with antigen (2,4-D). The anti- 2,4-D serum or monoclonal antibody was incubated with various amounts of 2,4-D. The solution was then drawn across the antigen-coated beads and anti-2,4-D antibodies with free Analyst, October 1997, Vol. 122 (1107–1111) 1107binding sites were bound to the beads. The amount of antibody bound to the beads was quantified by flowing a fluoresceinlabeled, species-specific antibody through the bead pack. This fluorescein-labeled antibody bound to the 2,4-D-specific antibody immobilized in the bead pack. The observed fluorescence was again inversely related to the concentration of analyte.The instrument flow system was operated under negative pressures using a syringe pump. An in-line vacuum de-gasser was used to prevent the formation of bubbles. All of the pumps and valves were operated through the KinExA hardware interface and PC-based software using a Windows environment. The instrument operations can be divided into two areas: bead handling and sample handling. For bead handling, the coated beads were drawn into the flow cell and removed to waste after the assay had been completed using a back-flush peristaltic pump.For sample handling, samples containing various concentrations of 2,4-D were drawn through channels 2–6 of a rotary valve, then over the packed bead column at a rate of 0.50 ml min21. The KinExA instrument software monitored the fluorescence signal once every second for a total of 420 s and displayed the data on the PC monitor. The fluorescence data were stored as voltage values in Excel and macros (included with the KinExA system) were used for analysis of the data.Signal intensities at specific time points in the instrument cycle were used as an indicator of the accumulation of fluorescent probe on the coated beads. The system was programmed to cycle automatically through the five samples, each using a new bead pack in the capillary. For the antibody-coated bead format, each cycle consisted of (1) bead packing, (2) analyte tracer accumulation and data acquisition, (3) buffer wash and (4) removal of the used beads to waste.For the antigen-coated bead format, step 2 was replaced by the accumulation of anti-2,4-D serum (incubated with various amounts of 2,4-D) followed by the accumulation of fluorescein-labeled anti-IgG antibody. Chemicals PMMA beads (98 mm) were purchased from Bang’s Laboratories (Carmel, IN, USA). Photopolymeric N-oxysuccinimde (PNOS) modified PMMA beads were purchased from BSI (Eden Prairie, MN, USA). Rabbit anti-sheep IgG and sheep anti-2,4-D-serum were obtained from The Binding Site (San Diego, CA, USA).Fluorescein-conjugated rabbit anti-sheep IgG was purchased from Jackson ImmunoResearch (West Grove, PA, USA). The anti-2,4-D monoclonal antibodies (clone E2/G2) were generated by Dr. Milan Franek at the Veterinary Research Institute (Brno, Czech Republic) and kindly provided by Dr. Sergei Erernin (Moscow State University, Russia). Fluorescein isothiocyanate (FITC), 2,4-D, 2,4,5-trichlorophenoxyacetic acid (2,4,5-T), l-ethyl-3-(3-dimethylaminopropyl)- carbodiimide hydrochloride (EDC), N-hydroxysuccimide and dimethylformamide were purchased from Sigma (St.Louis, MO, USA). Phenol and benzoic acid were purchased from Mallinckrodt (Paris, KY, USA). Atrazine was purchased from ChemService (West Chester, PA, USA). 2,4-Dichlorophenol was purchased from Aldrich (Milwaukee, WI, USA). Chlordane was obtained from the Environmental Protection Agency (Research Triangle Park, NC, USA).All other compounds and solvents used were of analytical-reagent grade. Environmental Samples River water samples were obtained from the Virgin River in Southern Utah. The water was passed through a coarse filter to remove any insoluble matter, and stored at 4 °C until use. The pH of the water at the time of use was 7.0. Well water was obtained from Well HM-1, Salmon site, Louisiana, it was stored at 4 °C until use, at which time its pH was 7.5. Preparation of Fluorescein-labeled 2,4-D Fluorescein thiocarbamylethylenediamine was synthesized from FITC and ethylenediamine as described by Pourfarzaneh et al.18 The amine-derivatized FITC was then conjugated to 2,4-D as reported by Eremin.19 Briefly, the N-hydroxysuccinimide ester of 2,4-D was synthesized and conjugated to fluorescein thiocarbamylethylenediamine in the presence of EDC in dimethylformamide.The tracer was purified by thinlayer chromatography [silica gel plates were developed in ethyl acetate–methanol–acetic acid (90 + 8 + 2, v/v/v)] and the concentration was determined spectrophotometrically using previously reported molar absorption coefficients.18 Preparation of Antibody-coated Beads Antibody-coated PMMA beads were prepared by immersing 200 mg of dry beads into 1 ml of anti-sheep IgG (20 mg ml21) in PBS (phosphate-buffered saline: 10 mm NaHPO4; 100 mm NaCl; pH 7.4).The mixture was then agitated for 2 h at 37 °C, after which the beads were centrifuged (5000g, 2 min) and the supernatant was discarded.The IgG-coated particles were rinsed twice by centrifugation with PBS (1 ml). The sheep anti- 2,4-D serum (diluted 1 + 199 into PBS) was immobilized onto the anti-sheep IgG-coated beads by incubation (with gentle rocking) for 2 h at 37 °C. The beads were again rinsed as described above, suspended in 1 ml of PBS and stored at 4 °C until the day of use, at which time they were diluted with 19 ml, of PBS and placed in the instrument reservoir.Preparation of Antigen-coated Beads Antigen-coated beads were prepared using amine reactive PNOS-coated beads. An amine derivative of 2,4-D was prepared by conjugating ethylenediamine to the N-hydroxysuccinimide ester of 2,4-D. The amine-derivatized 2,4-D was purified by thin-layer chromatography (silica gel plates developed in ethanol). Coupling of the ligand to the beads was carried out in 0.10 m borate buffer of pH 8.5, as suggested by the supplier (BSI). The mixture containing 200 mg of (PNOS) coated beads and 0.5 mm derivatized 2,4-D in 1 ml of the same buffer was agitated for 2 h at room temperature.The beads were then washed sequentially with 0.10 m borate buffer containing 1 m NaCl, followed by 0.10 m acetate buffer (pH 4) containing 1 m NaCl and finally with PBS. The beads were diluted with 19 ml of PBS for use in the instrument. Fig. 1 Diagrammatic representation of the immunoassay format for measurement of 2,4-D. (A) Competitive immunoassay format using antibody-coated beads, (B) competitive immunoassay format using antigencoated beads. 1108 Analyst, October 1997, Vol. 122Assay Procedure for Antibody-coated Beads 2,4-D stock solutions were prepared in ethanol and diluted to final concentrations as specified under Results and Discussion in PBS or an environmental water sample containing the fluorescein-labeled 2,4-D (1 nm or as specified under Results and Discussion). The final assay sample contained less than 1% ethanol which was included in the control and had no effect on the assay. The 420 s cycle (i.e., bead packing, tracer accumulation, buffer washing and bead expulsion) was then initiated through the KinExA software.Fluorescence signal data were collected, displayed and end-point measurements calculated from the difference in the average fluorescence signal of the initial five and last five data points. Assay Procedure for Antigen-coated Beads 2,4-D stock solutions were prepared in ethanol and diluted to final concentrations in PBS.Sheep anti-2,4-D serum diluted 1 + 39 999 in PBS or monoclonal antibody (E2/G2) at a final concentration of 12 ng ml21 was reacted with various concentrations of 2,4-D. The timing cycle was modified to include a step for the addition of the fluorescently labeled secondary antibody (either anti-rabbit or anti-mouse diluted 1 + 4999 in PBS containing 1 mg ml21 BSA). Fluorescence data were handled as above.Curve Fitting and Statistical Analysis The four parameter logistic equation as defined below was used to fit the immunoassay data.20 y a d x c d b = - + æ è ç ö ø ÷ + ( ) 1 where y = instrument response; a = response at high asymptote, b = slope factor; c = concentration corresponding to 50% specific binding (EC-50); d = response at low asymptote; and x = calibrator concentration. Calculations were performed using SOFTmax software (Molecular Devices, Menlo Park, CA, USA).Because the response error over the concentration range of the assay was heteroscedastic, an estimate of error for the precision of concentration was determined at a concentration near the EC-50 using the standard deviation/slope of the response curve at that point.20 Results and Discussion Shown in Fig. 2 are representative fluorescence signal tracings. There are three representative portions of the fluorescence tracing. First, the horizontal portion to the left shows the baseline fluorescence response for the packed beads prior to addition of the fluorescent tracer.Second, the steepest portion of the fluorescence curve represents the introduction of the bulk tracer into the capillary, filling the interstitial spaces between the beads (approximately the first 1–2 min), followed by the accumulation of the tracer on the coated bead surfaces. The third segment shows the removal of the free tracer from the bead pack. Tracings A, B and C represent control experiments for the antibody-coated bead format [shown in Fig. 1(a)]. Tracing A (Fig. 2) shows the non-specific signal when non-analyte specific sheep IgG was substituted for anti-2,4-D serum on the antibody-coated beads. Tracings B and C show the analytedependent response and result from the presence and absence, respectively, of 2,4-D (10 mg l21) in the assay buffer. The concentration-dependent signal response to 2,4-D was similar when determined from either the accumulation portion of the curve at 300 s or from the tracer remaining on the beads after a buffer wash measured by the fluorescence signal at 420 s.Readings at 420 s were routinely used to generate competition response curves. For most competitive immunoassay methods, the lowest potential detection limit of the assay is primarily determined by the affinity of the antibody for the antigen and, to a lesser extent, by the relative experimental error. However, for non-equilibrium methods such as is reported here, the relative concentrations of immobilized antibody and analyte probe may also impact the assay characteristics.Consequently, the effects of analyte tracer concentration and the polyclonal antibody immobilized on the solid phase were determined. For the antibody-coated bead format, analyte tracer concentrations were varied over the range 0.1–50 nm. A tracer concentration of 0.1 nm yielded no response to the analyte (2,4-D) and 50 nm tracer resulted in unacceptable levels of variability in the fluorescence response. A probe concentration of 1 nm yielded an acceptable signal and minimal variability.The antibody concentration used for immobilization was varied over the range of 0.1–2.0 mg protein per mg dry beads, and had no effect on the competition curve for 2,4-D (data not shown). Shown in Fig. 3 is the signal response curve for 2,4-D using the polyclonal antibody-coated bead format with fluoresceinlabeled 2,4-D as the fluorescent tracer.The mean fluorescence response data were fitted using a four parameter mode.20 The dynamic range extended from 10 mg l21 to 10 mg l21. The inflection point of the curve (EC-50) was 0.87 mg l21 and the multiple correlation coefficient (R) was 0.999. The precision of concentration was calculated at 1 ± 0.28 mg l21. Also shown in Fig. 3 is the signal response curve for 2,4-D, using the antigen-coated bead format for the polyclonal antiserum with fluorescein labeled anti-species antibody as the fluorescent tracer.The dynamic range for this format extended from 2 mg l21 to 20 mg l21. The EC-50 was 1.0 mg l21 and R was 0.998. The precision of concentration was calculated at 1 ± 0.12 mg l21. The competition curves for 2,4-D were similar for both assay formats suggesting that both assay formats were reporting the unoccupied (antibody) binding sites (in the presence of 2,4-D) and that the measuring process did not interfere with the steadystate association of the analyte with the antiserum.In addition, there are several advantages which may be realized by the versatility of this assay system. For example, the ability to link Fig. 2 Typical assay response and control experiments for antibodycoated bead format (A) non-specific binding signal; fluorescein-labeled 2,4-D (1 nm) was used with PMMA beads that were coated with nonanalyte- specific IgG. (B/C) IgG from anti-2,4-D serum was immobilized onto the PMMA beads (as described under Experimental).Antibody-coated beads were then exposed to the fluorescein-labeled 2,4-D (1 nm) in the presence (B) or absence (C) of target analyte 2,4-D (10 mg l21). Analyst, October 1997, Vol. 122 1109the antigen either to a fluorescent tag or to the derivatized beads (each with a different chemical linker) allows one to avoid similarities (and subsequent cross-reactivity) to the linker chain used to form the immunogen. This is particularly important for the use of commercially available antibodies for which the specific immunogen structure is not known.Shown in Fig. 4 are the signal response curves generated by spiking 2,4-D into various environmentally significant media, using polyclonal antiserum with the antibody-coated bead format. The signal response from the 2,4-D spiked well water significantly overlapped the signal response using buffer. The response for 2,4-D spiked into river water appeared slightly lower than that for the buffer or well water matrices.This bias was evidenced by the mean values for seven of the eight data points being lower than for the buffer control as well as the standard deviation values for four of the 2,4-D levels not overlapping those for the spiked buffer. The reason for the tendency of the river water matrix to result in an overestimation of the 2,4-D is not clear. 2,4-D contamination of the water was ruled out by HPLC analysis using standard methods.It is possible that compounds such as humic substances may have interfered with the assay. The relative errors for the environmental matrices were similar to those for the buffer control. The precision of concentration values calculated at l mg l21 were 1 ± 0.28 mg l21 for the buffer, 1 ± 0.27 mg l21 for the well water and 1 ± 0.43 mg l21 for the river water. Shown in Table 1 is the cross-reactivity profile for several structurally related compounds and structurally unrelated pesticides.As expected from previous reports of immunoassays using polyclonal anti-2,4-D serum, compounds similar in structure to 2,4-D, such as 2,4,5- and 2,4-dichlorophenol, elicit a response from this method.19 Compounds with structures unlike 2,4-D showed little if any response. The calibration plot for 2,4-D in Fig. 5 shows the response of the monoclonal antibody (E2/G2) using the antigen-coated bead format. The data were again fitted using the four parameter model.The EC-50 was 2.8 mg l21, R was 1.00 and the assay response curve showed a dynamic range of 0.1–100 mg l21. In comparison with the other reported bioanalytical methods for 2,4-D, the described KinExA method showed a higher response to the analyte than a reported enzyme assay,14 and fluorescence polarization immunoassay.13 The displacement curve and dynamic range for this method using the monoclonal antibody (E2/G2) are similar to those typically reported for ELISAs11 or ELISA-type assays4,17 and are about an order of magnitude less sensitive than microformat ELISAs using chemiluminescence detection.3 Nevertheless, given the profound influence of the affinity characteristics of any particular antibody on the assay response, comparison of immunoassay techniques that use different antibodies is tenuous.Conclusions A variety of immunoassay methods have been reported for the detection and measurement of 2,4-D, each with distinct analytical and assay format characteristics.The potential advantages of the proposed method in its application to environmental monitoring relates primarily to its ease of use (i.e., the degree of automation), the limited waste generated Fig. 3 Assay response as a function of 2,4-D concentration for the antibody-coated bead format (-), or the antigen-coated bead format (5). Error bars represent standard deviation, n = 3. Fig. 4 Effect of environmental water samples as assay matrix using the antibody-coated bead format. 2,4-D standard curves were prepared in PBS (-), spiked into well water (5) or river water (:) samples. Multiple regression coefficients were 0.999, 0.999 and 0.998 for the PBS, well water and river water, respectively. Error bars represent standard deviation, n = 3. Table 1 Cross-reactivity profile Compound Relative response* 2,4-D 100 2,4,5-T† 92 2,4-Dichlorophenol 86 Atrazine 20 Chlordane 14 Benzoic acid 10 Phenol 0 * Relative response to 2,4-D at 1 mg l21 determined using the following relationship: (response for test compound/response for 2,4-D) 3 100.† 2,4,5-Trichlorophenoxyacetic acid. Fig. 5 Assay response as a function of 2,4-D concentration using monoclonal antibody (E2/G2) in the antigen-coated bead format. 1110 Analyst, October 1997, Vol. 122(i.e., the absence of contaminated plates, tubes or vials), and the versatility allowed by the instrument and software in the development of immunoassays. The ability to use either the antibody or antigen as the immobilized phase allows the method to be tailored to the particular characteristics of the immunochemicals and requirements of the potential application (e.g., optimization of assay time and reagents per assay).The US Environmental Protection Agency (EPA), through its Office of Research and Development (ORD), funded the work involved in preparing this paper. It has been subject to the Agency’s peer review and has been approved for publication.Mention of trade names or commercial products does not constitute endorsement or recommendation by the US EPA. References 1 Meulenberg, E. P., Mulder, W. H., and Stoks, P.G., Environ. Sci. Technol., 1995, 29, 553. 2 Hall, J. C., Deschamps, R. J. A., and Krieg, K. K., J. Agric. Food Chem., 1989, 37, 981. 3 Dzgoev, A., Mecklenburg, M., Larson, P.-O., and Danielsson, B., Anal. Chem., 1996, 68, 3364. 4 Bauer, C. G., Eremenko, A. V., Ehrentreich-Forster, E., Bier, F. F., Makowwer, A., Halsall, H.B., Heineman, W. R., and Scheller, F. W., Anal. Chem., 1996, 68, 24S3. 5 Dohrman, C., Anal. Environ. Lab., 1991, October, 31. 6 Fleeker, J., J. Assoc. Off. Anal. Chem., 1994, 70, 874. 7 Ogert, R. A., Kusterbeck, A. W., Wemhoff, G. A., Burke, R., and Ligler, F. S., Anal. Lett., 1992, 25, 1999. 8 Alarie, J. P., Bowyer, J. R., Sepaniac, M. J., Hoyt, A. M., and Vo- Dinh, T., Anal. Chim. Acta, 1990, 236, 237. 9 Pollema, C. H., and Ruzicka, J., Anal. Chem., 1994, 66, 1825. 10 Mayer, M., and Ruzicka, J., Anal. Chem., 1996, 68, 3808. 11 Lawruk, T. S., Hottenstein, C. S., Fleeker, J. R., Hall, J. C., Herzog, D., and Rubio, F. M., Bull. Environ. Contam. Toxicol., 1994, 52, 538. 12 Lukin, Y. V., Dokuchaer, I. M., Polyak, I. M., and Eremin. S. A., Anal. Lett., 1994, 27, 2973. 13 Eremin, S. A., Landon, J., Smith, D. S., and Jackman, R., in Food Safety and Quality Assurance: Applications of Immunoassay Systems, ed. Morgan, M. R. A., Smith, C.J., and Williams, P. A., Elsevier Applied Science, New York, 1992. 14 Gu, Y., Knaebel, D. B., Korus, R. A., and Crawford, R. L., Environ. Sci. Technol., 1995, 29, 1622. 15 Suleiman, A. A., and Guilbault, G. G., Analyst, l994, 119, 2279. 16 Minunni, M., Skladal, P., and Mascini, M., Anal. Lett., 1994, 27, 1475. 17 Skladal, P., and Kalab, T., Anal. Chim. Acta, 1995, 316, 73. 18 Pourfarzaneh, M., White, G. W., Landon, J., and Smith, D. S., Clin. Chem., 1980, 26, 730. 19 Eremin, S.A., in Immunoanalysis of Agrochemicals, Emerging Technologies, ACS Symposium Series 586, ed. Nelson, J. O., Karu, A. E., and Wong, R. B., American Chemical Society, Washington, DC, 1995. 20 Howes, I., in Immunoassays Essential Data, ed. Edwards, R., Wiley, New York, 1996. Paper 7/01511I Received March 4, 1997 Accepted June 13, 1997 Analyst, October 1997, Vol. 122 1111 Detection of 2,4-Dichlorophenoxyacetic Acid Using a Fluorescence Immunoanalyzer Kim R. Rogers*a, Steven D.Kohla, Lee A. Riddicka and Thomas Glassb a US Environmental Protection Agency, National Exposure Research Laboratory, Las Vegas, NV 89193, USA b Sapidyne Inc., Boise, ID 83706, USA A flow immunoassay method for the measurement of 2,4-dichlorophenoxyacetic acid (2,4-D) was developed. The competitive fluorescence immunoassay relies on the use of antibody- or antigen-coated poly(methyl methacrylate) particles (98 mm diameter) as a renewable solid phase. The assay exhibits a dynamic range of 0.1–100 mg l21 using a monoclonal antibody or alternatively 10 mg l21 to 10 mg l21 using commercially available antiserum.The assay is demonstrated in buffered saline solution as well as in aquatic environmental media. The relative errors for the environmental matrices were similar to those for the buffer control. The precision of concentration values calculated at 1 mg l21 (for the assay using antiserum) were ±0.28, ±0.27 and ±0.43 mg l21 for the buffer, well water and river water matrices, respectively.The method shows cross-reactivity with compounds of closely related structure but little cross-reactivity with compounds dissimilar in structure to 2,4-D. The proposed automated competitive immunoassay method is rapid (between 7 and 15 min per assay), simple and potentially portable. Keywords: 2,4-Dichlorophenoxyacetic acid; fluoroimmunoassay; KinExA immunoanalyzer Methods for the rapid and cost-effective detection of pesticides in environmental settings have gained considerable attention in recent years, primarily due to concerns related to potential human exposure from spray run-off and spills into groundwater sources.1 Classical laboratory-based methods used to measure these compounds are typically time consuming and expensive.These methods usually require extensive extraction, derivatization and separation by GC or HPLC. To meet the need for faster and more cost-effective methods, a variety of techniques have recently been reported and a number of commercial products (methods) have been introduced.These methods are, for the most part, immunochemically based enzyme-linked immunosorbent assays (ELISA) using absorbance, fluorescence or electrochemical detection. These assays have typically been formatted for microtiter plates,2 thick film-based planar microwells, 3 microflow cells,4 test kits,5 or test tube-type optical spectrometers.6 Although automation has been incorporated into a number of these method formats, they typically require manual addition of reagents, several incubation steps, and physical manipulations such as changing tubes, trays or capillaries. Portable automated systems that avoid these physical manipulations and generate (as waste) only spent reagents show potential cost, time and waste disposal advantages for certain applications.Several approaches have been used for the automated immunoassay flow systems. Formats for these systems involve either regeneration of the antibody immobilized to the solid phase (i.e., disruption of antibody– antigen binding)7 or renewal of a mobile solid support (e.g., microspheres which are trapped in a flow cell) which has been coated with either the antibodies or antigens,8,9 or enzymes.10 The use of a mobile solid phase which is renewable for each assay offers certain technical advantages resulting in a fast, versatile and inexpensive assay system. 2,4-Dichlorophenoxyacetic acid (2,4-D) is widely used as a systemic herbicide for broad leaf weeds in a variety of crop and non-crop applications.This pesticide is typically monitored in environmental samples using GC methods which require extensive clean-up and derivatization, however, a number of potential field analytical methods have been reported. These methods include: ELISA3,4,9,11 enzyme immunoassay,1 immunoagglutination, 12 fluorescence polarization,13 and enzyme assays14 as well as several antibody-based biosensor methods. 15–17 Although most of these methods are portable and relatively simple to execute, they typically require multiple steps and generate solid wastes (e.g., plates, tubes and vials). In this paper, we describe the use of the KinExA immunoanalyzer to develop competitive fluorescence immunoassay methods for the detection of 2,4-D using both monoclonal and polyclonal antibodies. These methods use a mobile–renewable solid phase (i.e., microspheres) and can be fully automated.Experimental Instrumentation and KinExA Method The KinExA instrument is an automated fluorescence immunoassay system that uses microspheres as the solid phase (Sapidyne, Boise, ID, USA). For the assay formats described here, these beads [poly(methyl methacrylate), PMMA] were coated with either antibody or antigen and then packed into a capillary flow cell that is integrated into an epi-illumination filter fluorimeter system. For the format using antibody-coated beads, fluorescently labeled probe is introduced into the flow cell in the presence or absence of analyte and continuous interrogation of the bead column for analyte-sensitive accumulation of fluorescently labeled tracer on the solid phase forms the basis of this assay.For the format using antigen-coated beads, monoclonal antibody or anti-2,4-D antiserum is incubated with the analyte (2,4-D), then introduced into the flow cell containing antigencoated beads.This step is followed by fluorescently labeled secondary (species-specific, anti-IgG) antibody. For both formats, the observed fluorescence signal varies inversely with the analyte concentration. The competitive immunoassay format using the antibodycoated beads is described in Fig. 1(a). In this assay, the PMMA particles were coated with anti-sheep IgG which was used to capture and orient the primary antibody from sheep anti-2,4-D serum. Next, inhibition in the accumulation of fluoresceinlabeled 2,4-D tracer resulting from the presence of various concentrations of the analyte 2,4-D allowed the detection and quantification of this analyte. Another competitive immunoassay format described in Fig. 1(b) used beads coated with antigen (2,4-D). The anti- 2,4-D serum or monoclonal antibody was incubated with various amounts of 2,4-D. The solution was then drawn across the antigen-coated beads and anti-2,4-D antibodies with free Analyst, October 1997, Vol. 122 (1107–1111) 1107binding sites were bound to the beads.The amount of antibody bound to the beads was quantified by flowing a fluoresceinlabeled, species-specific antibody through the bead pack. This fluorescein-labeled antibody bound to the 2,4-D-specific antibody immobilized in the bead pack. The observed fluorescence was again inversely related to the concentration of analyte. The instrument flow system was operated under negative pressures using a syringe pump.An in-line vacuum de-gasser was used to prevent the formation of bubbles. All of the pumps and valves were operated through the KinExA hardware interface and PC-based software using a Windows environment. The instrument operations can be divided into two areas: bead handling and sample handling. For bead handling, the coated beads were drawn into the flow cell and removed to waste after the assay had been completed using a back-flush peristaltic pump. For sample handling, samples containing various concentrations of 2,4-D were drawn through channels 2–6 of a rotary valve, then over the packed bead column at a rate of 0.50 ml min21.The KinExA instrument software monitored the fluorescence signal once every second for a total of 420 s and displayed the data on the PC monitor. The fluorescence data were stored as voltage values in Excel and macros (included with the KinExA system) were used for analysis of the data. Signal intensities at specific time points in the instrument cycle were used as an indicator of the accumulation of fluorescent probe on the coated beads. The system was programmed to cycle automatically through the five samples, each using a new bead pack in the capillary.For the antibody-coated bead format, each cycle consisted of (1) bead packing, (2) analyte tracer accumulation and data acquisition, (3) buffer wash and (4) removal of the used beads to waste. For the antigen-coated bead format, step 2 was replaced by the accumulation of anti-2,4-D serum (incubated with various amounts of 2,4-D) followed by the accumulation of fluorescein-labeled anti-IgG antibody.Chemicals PMMA beads (98 mm) were purchased from Bang’s Laboratories (Carmel, IN, USA). Photopolymeric N-oxysuccinimde (PNOS) modified PMMA beads were purchased from BSI (Eden Prairie, MN, USA). Rabbit anti-sheep IgG and sheep anti-2,4-D-serum were obtained from The Binding Site (San Diego, CA, USA). Fluorescein-conjugated rabbit anti-sheep IgG was purchased from Jackson ImmunoResearch (West Grove, PA, USA). The anti-2,4-D monoclonal antibodies (clone E2/G2) were generated by Dr.Milan Franek at the Veterinary Research Institute (Brno, Czech Republic) and kindly provided by Dr. Sergei Erernin (Moscow State University, Russia). Fluorescein isothiocyanate (FITC), 2,4-D, 2,4,5-trichlorophenoxyacetic acid (2,4,5-T), l-ethyl-3-(3-dimethylaminopropyl)- carbodiimide hydrochloride (EDC), N-hydroxysuccimide and dimethylformamide were purchased from Sigma (St.Louis, MO, USA). Phenol and benzoic acid were purchased from Mallinckrodt (Paris, KY, USA). Atrazine was purchased from ChemService (West Chester, PA, USA). 2,4-Dichlorophenol was purchased from Aldrich (Milwaukee, WI, USA). Chlordane was obtained from the Environmental Protection Agency (Research Triangle Park, NC, USA). All other compounds and solvents used were of analytical-reagent grade. Environmental Samples River water samples were obtained from the Virgin River in Southern Utah.The water was passed through a coarse filter to remove any insoluble matter, and stored at 4 °C until use. The pH of the water at the time of use was 7.0. Well water was obtained from Well HM-1, Salmon site, Louisiana, it was stored at 4 °C until use, at which time its pH was 7.5. Preparation of Fluorescein-labeled 2,4-D Fluorescein thiocarbamylethylenediamine was synthesized from FITC and ethylenediamine as described by Pourfarzaneh et al.18 The amine-derivatized FITC was then conjugated to 2,4-D as reported by Eremin.19 Briefly, the N-hydroxysuccinimide ester of 2,4-D was synthesized and conjugated to fluorescein thiocarbamylethylenediamine in the presence of EDC in dimethylformamide. The tracer was purified by thinlayer chromatography [silica gel plates were developed in ethyl acetate–methanol–acetic acid (90 + 8 + 2, v/v/v)] and the concentration was determined spectrophotometrically using previously reported molar absorption coefficients.18 Preparation of Antibody-coated Beads Antibody-coated PMMA beads were prepared by immersing 200 mg of dry beads into 1 ml of anti-sheep IgG (20 mg ml21) in PBS (phosphate-buffered saline: 10 mm NaHPO4; 100 mm NaCl; pH 7.4).The mixture was then agitated for 2 h at 37 °C, after which the beads were centrifuged (5000g, 2 min) and the supernatant was discarded. The IgG-coated particles were rinsed twice by centrifugation with PBS (1 ml).The sheep anti- 2,4-D serum (diluted 1 + 199 into PBS) was immobilized onto the anti-sheep IgG-coated beads by incubation (with gentle rocking) for 2 h at 37 °C. The beads were again rinsed as described above, suspended in 1 ml of PBS and stored at 4 °C until the day of use, at which time they were diluted with 19 ml, of PBS and placed in the instrument reservoir. Preparation of Antigen-coated Beads Antigen-coated beads were prepared using amine reactive PNOS-coated beads.An amine derivative of 2,4-D was prepared by conjugating ethylenediamine to the N-hydroxysuccinimide ester of 2,4-D. The amine-derivatized 2,4-D was purified by thin-layer chromatography (silica gel plates developed in ethanol). Coupling of the ligand to the beads was carried out in 0.10 m borate buffer of pH 8.5, as suggested by the supplier (BSI). The mixture containing 200 mg of (PNOS) coated beads and 0.5 mm derivatized 2,4-D in 1 ml of the same buffer was agitated for 2 h at room temperature.The beads were then washed sequentially with 0.10 m borate buffer containing 1 m NaCl, followed by 0.10 m acetate buffer (pH 4) containing 1 m NaCl and finally with PBS. The beads were diluted with 19 ml of PBS for use in the instrument. Fig. 1 Diagrammatic representation of the immunoassay format for measurement of 2,4-D. (A) Competitive immunoassay format using antibody-coated beads, (B) competitive immunoassay format using antigencoated beads. 1108 Analyst, October 1997, Vol. 122Assay Procedure for Antibody-coated Beads 2,4-D stock solutions were prepared in ethanol and diluted to final concentrations as specified under Results and Discussion in PBS or an environmental water sample containing the fluorescein-labeled 2,4-D (1 nm or as specified under Results and Discussion). The final assay sample contained less than 1% ethanol which was included in the control and had no effect on the assay. The 420 s cycle (i.e., bead packing, tracer accumulation, buffer washing and bead expulsion) was then initiated through the KinExA software.Fluorescence signal data were collected, displayed and end-point measurements calculated from the difference in the average fluorescence signal of the initial five and last five data points. Assay Procedure for Antigen-coated Beads 2,4-D stock solutions were prepared in ethanol and diluted to final concentrations in PBS. Sheep anti-2,4-D serum diluted 1 + 39 999 in PBS or monoclonal antibody (E2/G2) at a final concentration of 12 ng ml21 was reacted with various concentrations of 2,4-D.The timing cycle was modified to include a step for the addition of the fluorescently labeled secondary antibody (either anti-rabbit or anti-mouse diluted 1 + 4999 in PBS containing 1 mg ml21 BSA). Fluorescence data were handled as above. Curve Fitting and Statistical Analysis The four parameter logistic equation as defined below was used to fit the immunoassay data.20 y a d x c d b = - + æ è ç ö ø ÷ + ( ) 1 where y = instrument response; a = response at high asymptote, b = slope factor; c = concentration corresponding to 50% specific binding (EC-50); d = response at low asymptote; and x = calibrator concentration.Calculations were performed using SOFTmax software (Molecular Devices, Menlo Park, CA, USA). Because the response error over the concentration range of the assay was heteroscedastic, an estimate of error for the precision of concentration was determined at a concentration near the EC-50 using the standard deviation/slope of the response curve at that point.20 Results and Discussion Shown in Fig. 2 are representative fluorescence signal tracings. There are three representative portions of the fluorescence tracing. First, the horizontal portion to the left shows the baseline fluorescence response for the packed beads prior to addition of the fluorescent tracer. Second, the steepest portion of the fluorescence curve represents the introduction of the bulk tracer into the capillary, filling the interstitial spaces between the beads (approximately the first 1–2 min), followed by the accumulation of the tracer on the coated bead surfaces.The third segment shows the removal of the free tracer from the bead pack. Tracings A, B and C represent control experiments for the antibody-coated bead format [shown in Fig. 1(a)]. Tracing A (Fig. 2) shows the non-specific signal when non-analyte specific sheep IgG was substituted for anti-2,4-D serum on the antibody-coated beads.Tracings B and C show the analytedependent response and result from the presence and absence, respectively, of 2,4-D (10 mg l21) in the assay buffer. The concentration-dependent signal response to 2,4-D was similar when determined from either the accumulation portion of the curve at 300 s or from the tracer remaining on the beads after a buffer wash measured by the fluorescence signal at 420 s.Readings at 420 s were routinely used to generate competition response curves. For most competitive immunoassay methods, the lowest potential detection limit of the assay is primarily determined by the affinity of the antibody for the antigen and, to a lesser extent, by the relative experimental error. However, for non-equilibrium methods such as is reported here, the relative concentrations of immobilized antibody and analyte probe may also impact the assay characteristics. Consequently, the effects of analyte tracer concentration and the polyclonal antibody immobilized on the solid phase were determined.For the antibody-coated bead format, analyte tracer concentrations were varied over the range 0.1–50 nm. A tracer concentration of 0.1 nm yielded no response to the analyte (2,4-D) and 50 nm tracer resulted in unacceptable levels of variability in the fluorescence response. A probe concentration of 1 nm yielded an acceptable signal and minimal variability.The antibody concentration used for immobilization was varied over the range of 0.1–2.0 mg protein per mg dry beads, and had no effect on the competition curve for 2,4-D (data not shown). Shown in Fig. 3 is the signal response curve for 2,4-D using the polyclonal antibody-coated bead format with fluoresceinlabeled 2,4-D as the fluorescent tracer. The mean fluorescence response data were fitted using a four parameter mode.20 The dynamic range extended from 10 mg l21 to 10 mg l21.The inflection point of the curve (EC-50) was 0.87 mg l21 and the multiple correlation coefficient (R) was 0.999. The precision of concentration was calculated at 1 ± 0.28 mg l21. Also shown in Fig. 3 is the signal response curve for 2,4-D, using the antigen-coated bead format for the polyclonal antiserum with fluorescein labeled anti-species antibody as the fluorescent tracer. The dynamic range for this format extended from 2 mg l21 to 20 mg l21.The EC-50 was 1.0 mg l21 and R was 0.998. The precision of concentration was calculated at 1 ± 0.12 mg l21. The competition curves for 2,4-D were similar for both assay formats suggesting that both assay formats were reporting the unoccupied (antibody) binding sites (in the presence of 2,4-D) and that the measuring process did not interfere with the steadystate association of the analyte with the antiserum. In addition, there are several advantages which may be realized by the versatility of this assay system.For example, the ability to link Fig. 2 Typical assay response and control experiments for antibodycoated bead format (A) non-specific binding signal; fluorescein-labeled 2,4-D (1 nm) was used with PMMA beads that were coated with nonanalyte- specific IgG. (B/C) IgG from anti-2,4-D serum was immobilized onto the PMMA beads (as described under Experimental). Antibody-coated beads were then exposed to the fluorescein-labeled 2,4-D (1 nm) in the presence (B) or absence (C) of target analyte 2,4-D (10 mg l21).Analyst, October 1997, Vol. 122 1109the antigen either to a fluorescent tag or to the derivatized beads (each with a different chemical linker) allows one to avoid similarities (and subsequent cross-reactivity) to the linker chain used to form the immunogen. This is particularly important for the use of commercially available antibodies for which the specific immunogen structure is not known.Shown in Fig. 4 are the signal response curves generated by spiking 2,4-D into various environmentally significant media, using polyclonal antiserum with the antibody-coated bead format. The signal response from the 2,4-D spiked well water significantly overlapped the signal response using buffer. The response for 2,4-D spiked into river water appeared slightly lower than that for the buffer or well water matrices. This bias was evidenced by the mean values for seven of the eight data points being lower than for the buffer control as well as the standard deviation values for four of the 2,4-D levels not overlapping those for the spiked buffer.The reason for the tendency of the river water matrix to result in an overestimation of the 2,4-D is not clear. 2,4-D contamination of the water was ruled out by HPLC analysis using standard methods. It is possible that compounds such as humic substances may have interfered with the assay.The relative errors for the environmental matrices were similar to those for the buffer control. The precision of concentration values calculated at l mg l21 were 1 ± 0.28 mg l21 for the buffer, 1 ± 0.27 mg l21 for the well water and 1 ± 0.43 mg l21 for the river water. Shown in Table 1 is the cross-reactivity profile for several structurally related compounds and structurally unrelated pesticides. As expected from previous reports of immunoassays using polyclonal anti-2,4-D serum, compounds similar in structure to 2,4-D, such as 2,4,5- and 2,4-dichlorophenol, elicit a response from this method.19 Compounds with structures unlike 2,4-D showed little if any response.The calibration plot for 2,4-D in Fig. 5 shows the response of the monoclonal antibody (E2/G2) using the antigen-coated bead format. The data were again fitted using the four parameter model. The EC-50 was 2.8 mg l21, R was 1.00 and the assay response curve showed a dynamic range of 0.1–100 mg l21.In comparison with the other reported bioanalytical methods for 2,4-D, the described KinExA method showed a higher response to the analyte than a reported enzyme assay,14 and fluorescence polarization immunoassay.13 The displacement curve and dynamic range for this method using the monoclonal antibody (E2/G2) are similar to those typically reported for ELISAs11 or ELISA-type assays4,17 and are about an order of magnitude less sensitive than microformat ELISAs using chemiluminescence detection.3 Nevertheless, given the profound influence of the affinity characteristics of any particular antibody on the assay response, comparison of immunoassay techniques that use different antibodies is tenuous.Conclusions A variety of immunoassay methods have been reported for the detection and measurement of 2,4-D, each with distinct analytical and assay format characteristics. The potential advantages of the proposed method in its application to environmental monitoring relates primarily to its ease of use (i.e., the degree of automation), the limited waste generated Fig. 3 Assay response as a function of 2,4-D concentration for the antibody-coated bead format (-), or the antigen-coated bead format (5). Error bars represent standard deviation, n = 3. Fig. 4 Effect of environmental water samples as assay matrix using the antibody-coated bead format. 2,4-D standard curves were prepared in PBS (-), spiked into well water (5) or river water (:) samples.Multiple regression coefficients were 0.999, 0.999 and 0.998 for the PBS, well water and river water, respectively. Error bars represent standard deviation, n = 3. Table 1 Cross-reactivity profile Compound Relative response* 2,4-D 100 2,4,5-T† 92 2,4-Dichlorophenol 86 Atrazine 20 Chlordane 14 Benzoic acid 10 Phenol 0 * Relative response to 2,4-D at 1 mg l21 determined using the following relationship: (response for test compound/response for 2,4-D) 3 100. † 2,4,5-Trichlorophenoxyacetic acid.Fig. 5 Assay response as a function of 2,4-D concentration using monoclonal antibody (E2/G2) in the antigen-coated bead format. 1110 Analyst, October 1997, Vol. 122(i.e., the absence of contaminated plates, tubes or vials), and the versatility allowed by the instrument and software in the development of immunoassays. The ability to use either the antibody or antigen as the immobilized phase allows the method to be tailored to the particular characteristics of the immunochemicals and requirements of the potential application (e.g., optimization of assay time and reagents per assay). The US Environmental Protection Agency (EPA), through its Office of Research and Development (ORD), funded the work involved in preparing this paper. It has been subject to the Agency’s peer review and has been approved for publication. Mention of trade names or commercial products does not constitute endorsement or recommendation by the US EPA. References 1 Meulenberg, E. P., Mulder, W. H., and Stoks, P.G., Environ. Sci. Technol., 1995, 29, 553. 2 Hall, J. C., Deschamps, R. J. A., and Krieg, K. K., J. Agric. Food Chem., 1989, 37, 981. 3 Dzgoev, A., Mecklenburg, M., Larson, P.-O., and Danielsson, B., Anal. Chem., 1996, 68, 3364. 4 Bauer, C. G., Eremenko, A. V., Ehrentreich-Forster, E., Bier, F. F., Makowwer, A., Halsall, H. B., Heineman, W. R., and Scheller, F. W., Anal. Chem., 1996, 68, 24S3. 5 Dohrman, C., Anal. Environ. Lab., 1991, October, 31. 6 Fleeker, J., J. Assoc. Off. Anal. Chem., 1994, 70, 874. 7 Ogert, R. A., Kusterbeck, A. W., Wemhoff, G. A., Burke, R., and Ligler, F. S., Anal. Lett., 1992, 25, 1999. 8 Alarie, J. P., Bowyer, J. R., Sepaniac, M. J., Hoyt, A. M., and Vo- Dinh, T., Anal. Chim. Acta, 1990, 236, 237. 9 Pollema, C. H., and Ruzicka, J., Anal. Chem., 1994, 66, 1825. 10 Mayer, M., and Ruzicka, J., Anal. Chem., 1996, 68, 3808. 11 Lawruk, T. S., Hottenstein, C. S., Fleeker, J. R., Hall, J. C., Herzog, D., and Rubio, F. M., Bull. Environ. Contam. Toxicol., 1994, 52, 538. 12 Lukin, Y. V., Dokuchaer, I. M., Polyak, I. M., and Eremin. S. A., Anal. Lett., 1994, 27, 2973. 13 Eremin, S. A., Landon, J., Smith, D. S., and Jackman, R., in Food Safety and Quality Assurance: Applications of Immunoassay Systems, ed. Morgan, M. R. A., Smith, C. J., and Williams, P. A., Elsevier Applied Science, New York, 1992. 14 Gu, Y., Knaebel, D. B., Korus, R. A., and Crawford, R. L., Environ. Sci. Technol., 1995, 29, 1622. 15 Suleiman, A. A., and Guilbault, G. G., Analyst, l994, 119, 2279. 16 Minunni, M., Skladal, P., and Mascini, M., Anal. Lett., 1994, 27, 1475. 17 Skladal, P., and Kalab, T., Anal. Chim. Acta, 1995, 316, 73. 18 Pourfarzaneh, M., White, G. W., Landon, J., and Smith, D. S., Clin. Chem., 1980, 26, 730. 19 Eremin, S. A., in Immunoanalysis of Agrochemicals, Emerging Technologies, ACS Symposium Series 586, ed. Nelson, J. O., Karu, A. E., and Wong, R. B., American Chemical Society, Washington, DC, 1995. 20 Howes, I., in Immunoassays Essential Data, ed. Edwards, R., Wiley, New York, 1996. Paper 7/01511I Received March 4, 1997 Accepted June 13, 1997 Analyst, October 1997, Vol. 122 1111

ISSN:0003-2654    

DOI:10.1039/a701511i

出版商:RSC    

年代:1997

数据来源: RSC

摘要:

Mixed Immunosorbent for Selective On-line Trace Enrichment and Liquid Chromatography of Phenylurea Herbicides in Environmental Waters A. Martin-Estebana, P. Fern�andeza, D. Stevensonb and C. C�amara*a a Departamento de Qu�ýmica Anal�ýtica, Facultad de Ciencias Qu�ýmicas, Universidad Complutense de Madrid, 28040 Madrid, Spain b Robens Institute, University of Surrey, Guildford, Surrey, UK GU2 5XH An immunosorbent containing antiisoproturon and antichlortoluron antibodies immobilised on aldehyde-activated silica was employed for the on-line preconcentration and liquid chromatography–diode array detection of several phenylureas.The efficiency of the coupling of the immunosorbent to the liquid chromatographic system and the characteristics of the immunosorbent were evaluated. The on-line system allowed the selective trace enrichment of chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron at the 0.05–0.5 mg l21 level in ground and river waters and provided detection limits in the range 0.01–0.03 mg l21 by percolating only 10 ml of water sample.The proposed method was validated by analysing freeze-dried tap water samples with a high content in pesticides of different chemical functionalities. The results obtained were compared with the mean value obtained in an interlaboratory exercise. Keywords: Phenylurea herbicides; immunosorbent; liquid chromatography; environmental waters Solid-phase extraction (SPE) is a powerful tool for sample handling in the analysis of water samples for pesticides.Moreover, the application of on-line coupling of SPE to liquid chromatography (LC)1 has allowed the determination of a wide variety of pesticides, reaching the detection limits required for the quality control of drinking water (0.1 mg l21 in Europe for a single pesticide).2 However, the lack of selectivity of the sorbents typically used (C18, apolar copolymers) prevents similar detection limits being achieved in surface water owing to the broad peak obtained at the beginning of the chromatogram and to the noisy baseline, which shows the need to search for highly selective sorbents.Immunosorbents have been used for years in medicine3 but only recently for pesticide determinations. An immunosorbent consists of antibodies against a target compound (antigen), immobilised on an appropriate support material. Theoretically, it would permit the selective preconcentration of the antigen (or related compounds) when a sample is run through the immunosorbent and subsequently eluted free of co-extractives.Once the analytes have been eluted, they can be determined by chromatographic techniques. In recent years, antibodies against atrazine,4 simazine,5 isoproturon4,6 and chlortoluron5,7 have been immobilised on aldehyde-activated silica and antibodies against atrazine8,9 have been immobilised on diol-activated silica, and they have been successfully employed to preconcentrate these pesticides and also closely related compounds able to bind to the antibody from environmental waters.The use of immunosorbents for the preconcentration of triazines,10 phenylurea herbicides11 and carbofuran12 in plant material has also been reported. In a previous study,13 the complementary action of antiisoproturon and antichlortoluron antibodies for the preconcentration of various phenylurea herbicides from environmental waters was demonstrated.The antibodies were successfully immobilised on aldehyde-activated silica and the mixed immunosorbent allowed the selective preconcentration of several phenylureas, which were subsequently eluted in 1 ml of a simple phosphate-buffered saline–ethanol mixture at pH 2. Since the immunosorbent can be reused and is pressure resistant, the aim of this work was to develop an on-line system coupled to LC for the determination of several phenylurea herbicides in environmental waters.This work was focused on the efficiency of coupling of the immunosorbent to the LC system, and on the parameters affecting analyte binding and desorption. The analytes selected were chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron. Experimental Apparatus Eluent delivery was provided by a ConstaMetric 4100 Series high pressure pump from Thermo Separation Products (Hemel Hempstead, Herts., UK) coupled with a SpectroMonitor 5000 photodiode-array detector from LDC Analytical (Riviera Beach, FL, USA).Stainless-steel precolumns (1 cm 3 4.6 mm id or 5 cm 3 4.6 mm id) were coupled to the loop of a Rheodyne (Cotati, CA, USA) Model 7725i injection valve in order to carry out the extraction and enrichment steps. The preconcentration pump was a Waters Model 590 from Millipore (Bedford, MA, USA). Reagents High-purity water from a Milli-Q system (Millipore) and RSgrade acetonitrile (Scharlau, Barcelona, Spain) were passed through a 0.45 mm nylon filter (Whatman, Maidstone, Kent, UK) before use.Chlortoluron, isoproturon, metobromuron, linuron, chlorbromuron and propanil were obtained from Riedel-de Ha�en (Hanover, Germany). Stock standard solutions (1000 mg l21) were prepared in acetonitrile and stored at 220 °C in the dark. Glacial acetic acid, disodium hydrogenorthophosphate, potassium dihydrogenorthophosphate, potassium chloride and sodium chloride were of analytical-reagent grade from Merck (Darmstadt, Germany) and Panreac (Barcelona, Spain).Phosphate-buffered saline (PBS) of pH 7.2–7.4 was prepared by adding 8.0 g of sodium chloride, 0.2 g of potassium chloride, 0.2 g of potassium dihydrogenorthophosphate and 2.9 g of disodium hydrogenorthophosphate to 1 l of Milli-Q-purified water. Preparation of the Mixed Immunosorbent Polyclonal antibodies against isoproturon and chlortoluron were raised in two different mature Suffolk sheep and collected Analyst, October 1997, Vol. 122 (1113–1117) 1113after a long period of immunization (66 weeks for antiisoproturon antibodies14 and 91 weeks for antichlortoluron antibodies15).The unpurified polyclonal antibodies against isoproturon and chlortoluron were separately immobilised on aldehyde-activated porous silica (particle size 90–130 mm, pore size 1000 Å) (Clifmar Associates, Guilford, UK) according to the following procedure. Disposable polypropylene separation columns (11.0 3 1.0 cm id; Lab M, No. D823), each provided with a polyethylene matrix support frit, were each packed with 0.5 g of aldehyde-activated silica and washed with 50 ml of PBS buffer.Next, 5 ml of PBS buffer were dispensed into each column followed by the addition of 100 ml of neat antiserum. The columns were then closed from both sides and left rolling on a rotamixer for 2 h at room temperature. Once again each column was washed with 10 ml of PBS buffer and 5 ml of 10 mm sodium cyanoborohydride (pH 6), made up with 1 m glycine buffer, was carefully added to the individual columns, which were then rotated overnight at room temperature. The next day, each column was washed with 10 ml of 0.3% HCl (pH 2) followed by 20 ml of PBS buffer.16 About 1 g of each immunosorbent was mixed to obtain an immunosorbent containing 2 g of dry solid phase and 200 ml of each antiserum.13 Stationary Phases and Columns The analytical column was a 25 cm 34.6 mm id column packed with Spherisorb 5 mm ODS2 from Symta (Madrid, Spain).The preconcentration step was carried out using a precolumn (1 cm 3 4.6 mm id) laboratory-packed with the styrene–divinylbenzene copolymer PRP-1 (Hamilton, Reno, NV, USA) or a precolumn (5 cm 3 4.6 mm id) filled manually with the mixed immunosorbent (approximately 0.35 g of dry material). Sample Preparation Ground and river water samples spiked with phenylureas at concentrations between 0.05 and 0.5 mg l21 were directly preconcentrated as explained below.Amounts of 0.5 g of freeze-dried tap water samples containing atrazine, simazine, carbaryl, propanil, linuron, fenamiphos and permethrin at concentration levels in the range 0.5–17.3 mg g21 were reconstituted by adding 0.5 l of 1023 mol l21 HCl according to the procedure recomended.17 Then 500 ml of the reconstituted water sample were diluted to 10 ml with PBS and preconcentrated on the immunosorbent or on ecolumns as explained below. Analytical Procedure Ground, river or reconstituted freeze-dried tap water samples (10 ml, pH 7) were filtered on a 0.45 mm nylon filter and then preconcentrated on the immunosorbent precolumn (also on the PRP-1 precolumn for the freeze-dried tap water samples) at a flow rate of 2 ml min21.After washing the precolumn with 10 ml of PBS, the valve was switched and the analytes were eluted and separated in the analytical column with the following elution gradient: from 65% A (0.005 mol l21 KH2PO4, pH 2 adjusted with glacial acetic acid) and 35% B (acetonitrile) to 25% A–75% B in 20 min and returning to the initial conditions in 10 min at a flow rate of 1 ml min21.The immunosorbent was conditioned before each analysis with 3 ml of PBS–ethanol (1 + 1) at pH 2 and 25 ml of PBS. The polymeric precolumn was conditioned with 10 ml of acetonitrile, 10 ml of water and finally with 10 ml of PBS. The immunosorbent was stored in PBS at 4 °C after use. Quantitative measurements of peak areas by LC–UV at 244 nm were carried out for all pesticides studied.Results and Discussion The pesticides selected in this study were chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron, in order to cover an intermediate degree of polarity. Based on previous experiments,13 more polar analytes (e.g., methoxuron) were discarded because they were weakly retained and had very low breakthrough volumes, and more hydrophobic analytes (e.g., chloroxuron) were ruled out because they were retained by the organic matrix dissolved in the sample, hindering their interaction with the antibodies and giving rise to low recoveries.On-line Immunoextraction Coupled to Liquid Chromatography There are several problems caused by coupling the immunosorbent to the LC system. Previous experiments13 demonstrated that more than 50 ng per gram of silica for each phenylurea herbicide in a mixture of all of them produced overloading of the immunosorbent and competition among the analytes for the free binding sites.Also, the presence of a large amount of organic solvent (ethanol) prevents on-line elution of the analytes from the immunosorbent and their re-preconcentration in a second precolumn packed with C18 or apolar copolymers that is subsequently coupled on-line to the LC system.8,9,12 Pichon et al.18 reported the direct coupling of an immunosorbent with antiisoproturon antibodies immobilised on aldehyde- activated silica to an LC system by using a long precolumn (3 cm 3 4.6 mm id), the analytes being eluted with an acetonitrile gradient.Theoretically, the peaks on the chromatogram should be broader than those obtained by direct injection; however, no band broadening was detected despite the large size of the precolumn. This may be because the retention in the immunosorbent is based on antigen–antibody interactions, which are not comparable to the hydrophobic interactions in the C18 analytical columns. With this in mind and since our immunosorbent had a very low capacity, a long precolumn (5 cm 3 4.6 mm id) was coupled directly to the loop of the injection valve and after the preconcentration step the analytes were eluted with an acetonitrile –water gradient.Although no band broadening was detected, it was not possible to elute the retained analytes completely. Quantitative recoveries and no memory effects were only obtained when the pH the mobile phase was < 3 and the injection valve was kept in the inject position up to a proportion of acetonitrile of 50%.The chromatogram obtained at 244 nm after preconcentration of 10 ml of Milli-Q-purified water spiked with 0.25 mg l21 of each phenylurea (Fig. 1) shows that the direct coupling of a long precolumn to the LC system is successfully achieved. It should be noted that the downward slope of the baseline on the chromatogram is due to the use of acetic acid, which absorbs at 244 nm and the content of which decreases owing to the applied gradient. Capacity of the Immunosorbent The capacity of the immunosorbent depends on the total number of sites available to bind the antigen.The immunosorbent used in this study was prepared with unpurified polyclonal antibodies against isoproturon and chlortoluron and thus contained at least two different kinds of antibodies. This implies that the interaction (and therefore the capacity of the immunosorbent) of a given analyte with the immunosorbent will be different depending on whether it is alone or in a mixture of related compounds.To carry out this study, the immunosorbent was loaded with 10 ml of Milli-Q-purified water spiked with the herbicides to be tested at a concentration level within the range 0.025–12 mg l21 when each phenylurea was alone in the sample or within the 1114 Analyst, October 1997, Vol. 122range 0.025–4 mg l21 when they were all together in the mixture. Fig. 2 shows the binding curves obtained, which are plots of the amount of each herbicide loaded on the immunosorbent versus the concentration in the percolated sample when the herbicides were (A) alone or (B) in a mixture. The slopes obtained for each herbicide in the linear region [Fig. 2(A)] show that all the pesticides tested have the same affinity, while the capacity varies from 20 ng for isoproturon to > 50 ng for chlorbromuron. This may be because when unspecific antibodies are used, the differential recognition of similar compounds is based on molecular mass and/or hydrophobicity. 19 The binding curves for isoproturon and chlortoluron show two different linear regions, which may indicate the presence of different kinds of antibodies in the immunosorbent, as confirmed by the results in Fig. 2(B). It is clear that the binding curves for isoproturon and chlortoluron reach a plateau when the total amount of herbicides is high, which indicates the presence of specific antibodies against isoproturon and chlortoluron and it is possible to load around 5 ng for isoproturon and 10 ng for chlortoluron when they are in a mixture because the competition will take place among the other herbicides present in the sample.This is clearly illustrated by the shape of the metobromuron binding curve, which shows a decrease in the amount loaded when the concentration of herbicides is high and, as indicated above, recognition depends on the molecular mass and polarity and so the analytes compete for the free binding sites.In summary, the immunosorbent used in this study contained three different kinds of antibodies: one specific for isoproturon, another specific for chlortoluron and the third consisting of unspecific antibodies. The results of this study suggest that it should be possible to preconcentrate the phenylurea herbicides tested at concentrations < 0.5 mg l21 when 10 ml of sample are percolated. Although this capacity is very low, it should allow the determination of the selected analytes at the low levels present in real water samples, because of the high selectivity of the antigen–antibody interactions.Binding Flow Rate The binding flow rate was evaluated within the range 0.5–3 ml min21 by preconcentration of 10 ml of Milli-Q-purified water spiked with herbicides at 0.25 mg l21. The only effect detected was a decrease of 50% in the efficiency of preconcentration for metobromuron when the sample flow rate was increased from 2 to 3 ml min21.The recoveries of the other phenylureas were not affected by increasing the sample flow rate. In order to obtain quantitative recoveries for all the herbicides studied, a flow rate of 2 ml min21 was chosen as the optimum. Breakthrough Volumes The effect of sample volume was studied at a constant mass of 2.5 ng of each phenylurea using 10, 25 and 50 ml of Milli-Qpurified water fortified with 0.25, 0.1 and 0.05 mg l21 of each analyte, respectively. The results obtained show that the sample volume (within the range studied) did not affect the preconcentration of the selected pesticides, except for metobromuron, for which a 20% decrease in preconcentration efficiency was observed when the sample volume was 50 ml.Selectivity: Application to Different Matrices The efficiency of the immunosorbent for the selective preconcentration of chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron from ground, river and freeze-dried tap water samples was evaluated.Fig. 3 shows the chromatograms obtained from on-line preconcentration on the immunosorbent of 10 ml of (A) ground water and (B) river water spiked at the 0.05 mg l21 level with each phenylurea. It clearly demonstrates the high degree of clean-up obtained with the proposed method, since the baselines in the two chromatograms are comparable and as clean as that obtained by preconcentrating Milli-Q-purified water (see Fig. 1). This should allow calibration graphs to be constructed for all analytes in Milli-Q-purified water and to be used for any kind of water. Freeze-dried tap water sample containing pesticides with different chemical functionalities [atrazine and simazine (tria- Fig. 1 LC–UV trace obtained at 244 nm after on-line immunoextraction of 10 ml of Milli-Q-purified water spiked with 0.25 mg l21 of each phenylurea herbicide. Peaks: 1 = chlortoluron, 2 = isoproturon; 3 = metobromuron; 4 = linuron and 5 = chlorbromuron.The asterisk indicates an impurity arising from the synthesis of the immunosorbent. For LC conditions, see Experimental. Fig. 2 Binding curves for chlortoluron (1), isoproturon (2), metobromuron (3), linuron (4) and chlorbromuron (5) after on-line immunoextraction of 10 ml of Milli-Q-purified water sample when the compounds are alone (A) or mixed (B). See text for details. Analyst, October 1997, Vol. 122 1115zines), carbaryl (carbamate), propanil (propioanilide), linuron (phenylurea), fenamiphos (organophosphorus) and permethrin (pyrethroid)] at concentrations within the range 0.5–17.3 mg g21 was also preconcentrated in order to demonstrate the high selectivity of the immunosorbent.Fig. 4 shows the chromatograms obtained after on-line preconcentration of 10 ml of diluted reconstituted tap water sample (see Experimental), (A) on a precolumn packed with the apolar copolymer PRP-1 and (B) on the immunosorbent.The chromatogram is cleaner after preconcentration on the immunosorbent than on the PRP-1 precolumn, because the antibodies immobilised are only able to retain phenylurea herbicides. However, in Fig. 4(B), two peaks are observed, one at the retention time of linuron and another which was identified as propanil by constructing its specific calibration graph in Milli-Q-purified water using the procedure described. Although propanil belongs to another family of pesticides (propioanilides), there is a clear similarity between the structures of propanil and linuron, as shown in Fig. 4(B), and the antibodies are not able to differentiate between compounds with this high degree of structural similarity. This would make it possible to increase the number of compounds that the immunosorbent is able to retain, such as anilines, which are transformation products from the phenylurea herbicides and some of which are included in the list of priority pollutants to be monitored in environmental waters in Europe.This interesting possibility is now under study. Analytical Performance and Method Validation Calibration graphs for ground and river water were constructed by applying the on-line immunoextraction procedure to 10 ml of spiked water within the range 0.05–0.5 mg l21 and the resulting correlation coefficients were satisfactory (r2 > 0.99). The RSD for the selected analytes at a concentration of 0.25 mg l21 using the full procedure and evaluating the peak areas was in the range 1–9% (n = 5), depending on the pesticide.The detection limits, which were calculated as three times the standard deviation of the lowest concentration solutions, were within the range 0.01–0.03 mg l21, depending on the phenylurea herbicide, although they could be lowered further by preconcentrating a larger sample volume. These low detection limits obtained with only 10 ml of sample are a remarkable result and are good enough to allow the fate and transport of phenylureas to be studied directly in environmental waters.Calibration graphs were also constructed for propanil and linuron within the range 0.05–0.5 mg l21 by the on-line immunoextraction of 10 ml of Milli-Q-purified water and were used for the determination of propanil and linuron in the freezedried tap water samples. The results (Table 1) were in good agreement with the mean values obtained in an interlaboratory exercise,17 in which only validated methods were used. Reusability of the Immunosorbent The immunosorbent used in this study was employed in the analysis of more than 50 samples, including dirty water samples such as river water, and no decrease in the capacity of the immunosorbent was detected.This long lifetime is due to the good stability of the antibodies immobilised on aldehyde- Fig. 3 LC–UV traces obtained at 244 nm after on-line immunoextraction of 10 ml of ground (A) and river (B) water spiked with 0.05 mg l21 of each phenylurea herbicide.Peak numbers as in Fig. 1. For LC conditions, see Experimental. Fig. 4 LC–UV traces obtained at 244 nm after on-line preconcentration of 10 ml of freeze-dried reconstituted tap water sample on a PRP-1 precolumn (A) and on the immunosorbent (B). For LC conditions, see Experimental. Table 1 Mean values ± standard deviations (mg g21) obtained in the determination of propanil and linuron in freeze-dried tap water samples Interlaboratory On-line Compound exercise* immunoextraction† Propanil 11.70 ± 1.31 11.12 ± 0.24 Linuron 5.47 ± 0.90 5.66 ± 0.36 * Data obtained from ref. 17. † Mean value of three independent determinations. 1116 Analyst, October 1997, Vol. 122activated silica and because the regeneration (25 ml of PBS) and storage were appropriate for the period tested. Conclusion The use of a precolumn containing a mixed immunosorbent with immobilised antiisoproturon and antichlortoluron antibodies allows the simultaneous on-line immunopreconcentration of several phenylurea herbicides from ground and river water.The high selectivity shown by the immunosorbent allows the determination of phenylurea herbicides at trace levels when only 10 ml of water sample are percolated. Moreover, the selectivity of the immunosorbent was clearly demonstrated by analysing freeze-dried tap water samples spiked with large amounts of pesticides with different chemical functionalities, and the immunosorbent only retained linuron (phenylurea herbicide) and propanil (propioanilide), which show a high degree of structural similarity.Although this immunosorbent is not specific for isoproturon and chlortoluron, it offers the advantage of recognizing the phenylurea family of herbicides and very closely related compounds, which may be of great interest for environmental analysis. This work represents yet another example of the great potential shown by the immunosorbents used to date for the preconcentration of pesticides and further development in this area should be undertaken.This work received financial support from the Standards, Measurements and Testing Programme under contract number MAT1-CT940001 and PB95-0366-C02-01. The authors thank Max Gorman for revision of the manuscript. References 1 Barcel�o, D., and Hennion, M. C., Anal. Chim. Acta, 1995, 318, 1. 2 EC Directive relating quality of water intended for human consumption (80/778/EC), Off.J. Eur. Communities, L229/11, 1980. 3 van Ginkel, L. A., Stephany, R. W., van Rossum, H. J., and Zoontjes, P. W., Trends Anal. Chem., 1992, 11, 294. 4 Pichon, V., Chen, L., Hennion, M. C., Daniel, R., Martel, A., Le Goffic, F., Abian, J., and Barcel�o, D., Anal. Chem., 1995, 67, 2451. 5 Pichon, V., Chen, L., Durand, N., Le Goffic, F., and Hennion, M. C., J. Chromatogr. A, 1996, 725, 107. 6 Shahtaheri, S. J., Kwasowski, P., and Stevenson, D., J. Chromatogr.A, submitted for publication. 7 Shahtaheri, S. J., Katmeh, M .F., Kwasowski, P., and Stevenson, D., J. Chromatogr. A, 1995, 697, 131. 8 Thomas, D. H., Beck-Westermeyer, M., and Hage, D. S., Anal. Chem., 1994, 66, 3823. 9 Rollag, J. G., Beck-Westermeyer, M., and Hage, D. S., Anal. Chem., 1996, 68, 3631. 10 Lawrence, J. Fenard, C., Hennion, M. C., Pichon, V., Le Goffic, F., and Durand, N., J. Chromatogr. A, 1996, 752, 147. 11 Lawrence, J. F., Menard, C., Hennion, M.C., Pichon, V., Le Goffic, F., and Durand, N., J. Chromatogr. A, 1996, 732, 277. 12 Rule, G. S., Mordehai, A. V., and Henion, J., Anal. Chem., 1994, 66, 230. 13 Mart�ýn-Esteban, A., Kwasowski, P., and Stevenson, D., Chromatographia, 1997, 45, 364. 14 Katmeh, M. F., Frost, G., Aherne, W., and Stevenson, D., Analyst, 1994, 119, 431. 15 Katmeh, M. F., Aherne, W., and Stevenson, D., Analyst, 1996, 121, 1699. 16 Katmeh, M. F., Thesis, Robens Institute, University of Surrey, Guidford, 1994. 17 Mart�ýn-Esteban, A., Fern�andez, P., C�amara, C., Kramer, G. N., and Maier, E. A., Int. J. Environ. Anal. Chem., in the press. 18 Pichon, V., Chen, L., and Hennion, M. C., Anal. Chim. Acta, 1995, 311, 429. 19 Price, C. P., and Newman, D. J., Principles and Practice of Immunoassay, Macmillan, London, 1991. Paper 7/02828H Received April 25, 1997 Accepted July 22, 1997 Analyst, October 1997, Vol. 122 1117 Mixed Immunosorbent for Selective On-line Trace Enrichment and Liquid Chromatography of Phenylurea Herbicides in Environmental Waters A.Martin-Estebana, P. Fern�andeza, D. Stevensonb and C. C�amara*a a Departamento de Qu�ýmica Anal�ýtica, Facultad de Ciencias Qu�ýmicas, Universidad Complutense de Madrid, 28040 Madrid, Spain b Robens Institute, University of Surrey, Guildford, Surrey, UK GU2 5XH An immunosorbent containing antiisoproturon and antichlortoluron antibodies immobilised on aldehyde-activated silica was employed for the on-line preconcentration and liquid chromatography–diode array detection of several phenylureas.The efficiency of the coupling of the immunosorbent to the liquid chromatographic system and the characteristics of the immunosorbent were evaluated. The on-line system allowed the selective trace enrichment of chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron at the 0.05–0.5 mg l21 level in ground and river waters and provided detection limits in the range 0.01–0.03 mg l21 by percolating only 10 ml of water sample.The proposed method was validated by analysing freeze-dried tap water samples with a high content in pesticides of different chemical functionalities. The results obtained were compared with the mean value obtained in an interlaboratory exercise. Keywords: Phenylurea herbicides; immunosorbent; liquid chromatography; environmental waters Solid-phase extraction (SPE) is a powerful tool for sample handling in the analysis of water samples for pesticides. Moreover, the application of on-line coupling of SPE to liquid chromatography (LC)1 has allowed the determination of a wide variety of pesticides, reaching the detection limits required for the quality control of drinking water (0.1 mg l21 in Europe for a single pesticide).2 However, the lack of selectivity of the sorbents typically used (C18, apolar copolymers) prevents similar detection limits being achieved in surface water owing to the broad peak obtained at the beginning of the chromatogram and to the noisy baseline, which shows the need to search for highly selective sorbents.Immunosorbents have been used for years in medicine3 but only recently for pesticide determinations. An immunosorbent consists of antibodies against a target compound (antigen), immobilised on an appropriate support material. Theoretically, it would permit the selective preconcentration of the antigen (or related compounds) when a sample is run through the immunosorbent and subsequently eluted free of co-extractives. Once the analytes have been eluted, they can be determined by chromatographic techniques.In recent years, antibodies against atrazine,4 simazine,5 isoproturon4,6 and chlortoluron5,7 have been immobilised on aldehyde-activated silica and antibodies against atrazine8,9 have been immobilised on diol-activated silica, and they have been successfully employed to preconcentrate these pesticides and also closely related compounds able to bind to the antibody from environmental waters.The use of immunosorbents for the preconcentration of triazines,10 phenylurea herbicides11 and carbofuran12 in plant material has also been reported. In a previous study,13 the complementary action of antiisoproturon and antichlortoluron antibodies for the preconcentration of various phenylurea herbicides from environmental waters was demonstrated. The antibodies were successfully immobilised on aldehyde-activated silica and the mixed immunosorbent allowed the selective preconcentration of several phenylureas, which were subsequently eluted in 1 ml of a simple phosphate-buffered saline–ethanol mixture at pH 2.Since the immunosorbent can be reused and is pressure resistant, the aim of this work was to develop an on-line system coupled to LC for the determination of several phenylurea herbicides in environmental waters. This work was focused on the efficiency of coupling of the immunosorbent to the LC system, and on the parameters affecting analyte binding and desorption.The analytes selected were chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron. Experimental Apparatus Eluent delivery was provided by a ConstaMetric 4100 Series high pressure pump from Thermo Separation Products (Hemel Hempstead, Herts., UK) coupled with a SpectroMonitor 5000 photodiode-array detector from LDC Analytical (Riviera Beach, FL, USA). Stainless-steel precolumns (1 cm 3 4.6 mm id or 5 cm 3 4.6 mm id) were coupled to the loop of a Rheodyne (Cotati, CA, USA) Model 7725i injection valve in order to carry out the extraction and enrichment steps.The preconcentration pump was a Waters Model 590 from Millipore (Bedford, MA, USA). Reagents High-purity water from a Milli-Q system (Millipore) and RSgrade acetonitrile (Scharlau, Barcelona, Spain) were passed through a 0.45 mm nylon filter (Whatman, Maidstone, Kent, UK) before use.Chlortoluron, isoproturon, metobromuron, linuron, chlorbromuron and propanil were obtained from Riedel-de Ha�en (Hanover, Germany). Stock standard solutions (1000 mg l21) were prepared in acetonitrile and stored at 220 °C in the dark. Glacial acetic acid, disodium hydrogenorthophosphate, potassium dihydrogenorthophosphate, potassium chloride and sodium chloride were of analytical-reagent grade from Merck (Darmstadt, Germany) and Panreac (Barcelona, Spain). Phosphate-buffered saline (PBS) of pH 7.2–7.4 was prepared by adding 8.0 g of sodium chloride, 0.2 g of potassium chloride, 0.2 g of potassium dihydrogenorthophosphate and 2.9 g of disodium hydrogenorthophosphate to 1 l of Milli-Q-purified water.Preparation of the Mixed Immunosorbent Polyclonal antibodies against isoproturon and chlortoluron were raised in two different mature Suffolk sheep and collected Analyst, October 1997, Vol. 122 (1113–1117) 1113after a long period of immunization (66 weeks for antiisoproturon antibodies14 and 91 weeks for antichlortoluron antibodies15).The unpurified polyclonal antibodies against isoproturon and chlortoluron were separately immobilised on aldehyde-activated porous silica (particle size 90–130 mm, pore size 1000 Å) (Clifmar Associates, Guilford, UK) according to the following procedure. Disposable polypropylene separation columns (11.0 3 1.0 cm id; Lab M, No. D823), each provided with a polyethylene matrix support frit, were each packed with 0.5 g of aldehyde-activated silica and washed with 50 ml of PBS buffer.Next, 5 ml of PBS buffer were dispensed into each column followed by the addition of 100 ml of neat antiserum. The columns were then closed from both sides and left rolling on a rotamixer for 2 h at room temperature. Once again each column was washed with 10 ml of PBS buffer and 5 ml of 10 mm sodium cyanoborohydride (pH 6), made up with 1 m glycine buffer, was carefully added to the individual columns, which were then rotated overnight at room temperature.The next day, each column was washed with 10 ml of 0.3% HCl (pH 2) followed by 20 ml of PBS buffer.16 About 1 g of each immunosorbent was mixed to obtain an immunosorbent containing 2 g of d00 ml of each antiserum.13 Stationary Phases and Columns The analytical column was a 25 cm 34.6 mm id column packed with Spherisorb 5 mm ODS2 from Symta (Madrid, Spain). The preconcentration step was carried out using a precolumn (1 cm 3 4.6 mm id) laboratory-packed with the styrene–divinylbenzene copolymer PRP-1 (Hamilton, Reno, NV, USA) or a precolumn (5 cm 3 4.6 mm id) filled manually with the mixed immunosorbent (approximately 0.35 g of dry material). Sample Preparation Ground and river water samples spiked with phenylureas at concentrations between 0.05 and 0.5 mg l21 were directly preconcentrated as explained below.Amounts of 0.5 g of freeze-dried tap water samples containing atrazine, simazine, carbaryl, propanil, linuron, fenamiphos and permethrin at concentration levels in the range 0.5–17.3 mg g21 were reconstituted by adding 0.5 l of 1023 mol l21 HCl according to the procedure recomended.17 Then 500 ml of the reconstituted water sample were diluted to 10 ml with PBS and preconcentrated on the immunosorbent or on the PRP-1 precolumns as explained below. Analytical Procedure Ground, river or reconstituted freeze-dried tap water samples (10 ml, pH 7) were filtered on a 0.45 mm nylon filter and then preconcentrated on the immunosorbent precolumn (also on the PRP-1 precolumn for the freeze-dried tap water samples) at a flow rate of 2 ml min21.After washing the precolumn with 10 ml of PBS, the valve was switched and the analytes were eluted and separated in the analytical column with the following elution gradient: from 65% A (0.005 mol l21 KH2PO4, pH 2 adjusted with glacial acetic acid) and 35% B (acetonitrile) to 25% A–75% B in 20 min and returning to the initial conditions in 10 min at a flow rate of 1 ml min21.The immunosorbent was conditioned before each analysis with 3 ml of PBS–ethanol (1 + 1) at pH 2 and 25 ml of PBS. The polymeric precolumn was conditioned with 10 ml of acetonitrile, 10 ml of water and finally with 10 ml of PBS. The immunosorbent was stored in PBS at 4 °C after use. Quantitative measurements of peak areas by LC–UV at 244 nm were carried out for all pesticides studied.Results and Discussion The pesticides selected in this study were chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron, in order to cover an intermediate degree of polarity. Based on previous experiments,13 more polar analytes (e.g., methoxuron) were discarded because they were weakly retained and had very low breakthrough volumes, and more hydrophobic analytes (e.g., chloroxuron) were ruled out because they were retained by the organic matrix dissolved in the sample, hindering their interaction with the antibodies and giving rise to low recoveries.On-line Immunoextraction Coupled to Liquid Chromatography There are several problems caused by coupling the immunosorbent to the LC system. Previous experiments13 demonstrated that more than 50 ng per gram of silica for each phenylurea herbicide in a mixture of all of them produced overloading of the immunosorbent and competition among the analytes for the free binding sites.Also, the presence of a large amount of organic solvent (ethanol) prevents on-line elution of the analytes from the immunosorbent and their re-preconcentration in a second precolumn packed with C18 or apolar copolymers that is subsequently coupled on-line to the LC system.8,9,12 Pichon et al.18 reported the direct coupling of an immunosorbent with antiisoproturon antibodies immobilised on aldehyde- activated silica to an LC system by using a long precolumn (3 cm 3 4.6 mm id), the analytes being eluted with an acetonitrile gradient.Theoretically, the peaks on the chromatogram should be broader than those obtained by direct injection; however, no band broadening was detected despite the large size of the precolumn. This may be because the retention in the immunosorbent is based on antigen–antibody interactions, which are not comparable to the hydrophobic interactions in the C18 analytical columns. With this in mind and since our immunosorbent had a very low capacity, a long precolumn (5 cm 3 4.6 mm id) was coupled directly to the loop of the injection valve and after the preconcentration step the analytes were eluted with an acetonitrile –water gradient.Although no band broadening was detected, it was not possible to elute the retained analytes completely. Quantitative recoveries and no memory effects were only obtained when the pH the mobile phase was < 3 and the injection valve was kept in the inject position up to a proportion of acetonitrile of 50%.The chromatogram obtained at 244 nm after preconcentration of 10 ml of Milli-Q-purified water spiked with 0.25 mg l21 of each phenylurea (Fig. 1) shows that the direct coupling of a long precolumn to the LC system is successfully achieved. It should be noted that the downward slope of the baseline on the chromatogram is due to the use of acetic acid, which absorbs at 244 nm and the content of which decreases owing to the applied gradient.Capacity of the Immunosorbent The capacity of the immunosorbent depends on the total number of sites available to bind the antigen. The immunosorbent used in this study was prepared with unpurified polyclonal antibodies against isoproturon and chlortoluron and thus contained at least two different kinds of antibodies. This implies that the interaction (and therefore the capacity of the immunosorbent) of a given analyte with the immunosorbent will be different depending on whether it is alone or in a mixture of related compounds.To carry out this study, the immunosorbent was loaded with 10 ml of Milli-Q-purified water spiked with the herbicides to be tested at a concentration level within the range 0.025–12 mg l21 when each phenylurea was alone in the sample or within the 1114 Analyst, October 1997, Vol. 122range 0.025–4 mg l21 when they were all together in the mixture. Fig. 2 shows the binding curves obtained, which are plots of the amount of each herbicide loaded on the immunosorbent versus the concentration in the percolated sample when the herbicides were (A) alone or (B) in a mixture.The slopes obtained for each herbicide in the linear region [Fig. 2(A)] show that all the pesticides tested have the same affinity, while the capacity varies from 20 ng for isoproturon to > 50 ng for chlorbromuron. This may be because when unspecific antibodies are used, the differential recognition of similar compounds is based on molecular mass and/or hydrophobicity. 19 The binding curves for isoproturon and chlortoluron show two different linear regions, which may indicate the presence of different kinds of antibodies in the immunosorbent, as confirmed by the results in Fig. 2(B). It is clear that the binding curves for isoproturon and chlortoluron reach a plateau when the total amount of herbicides is high, which indicates the presence of specific antibodies against isoproturon and chlortoluron and it is possible to load around 5 ng for isoproturon and 10 ng for chlortoluron when they are in a mixture because the competition will take place among the other herbicides present in the sample.This is clearly illustrated by the shape of the metobromuron binding curve, which shows a decrease in the amount loaded when the concentration of herbicides is high and, as indicated above, recognition depends on the molecular mass and polarity and so the analytes compete for the free binding sites.In summary, the immunosorbent used in this study contained three different kinds of antibodies: one specific for isoproturon, another specific for chlortoluron and the third consisting of unspecific antibodies. The results of this study suggest that it should be possible to preconcentrate the phenylurea herbicides tested at concentrations < 0.5 mg l21 when 10 ml of sample are percolated. Although this capacity is very low, it should allow the determination of the selected analytes at the low levels present in real water samples, because of the high selectivity of the antigen–antibody interactions.Binding Flow Rate The binding flow rate was evaluated within the range 0.5–3 ml min21 by preconcentration of 10 ml of Milli-Q-purified water spiked with herbicides at 0.25 mg l21. The only effect detected was a decrease of 50% in the efficiency of preconcentration for metobromuron when the sample flow rate was increased from 2 to 3 ml min21.The recoveries of the other phenylureas were not affected by increasing the sample flow rate. In order to obtain quantitative recoveries for all the herbicides studied, a flow rate of 2 ml min21 was chosen as the optimum. Breakthrough Volumes The effect of sample volume was studied at a constant mass of 2.5 ng of each phenylurea using 10, 25 and 50 ml of Milli-Qpurified water fortified with 0.25, 0.1 and 0.05 mg l21 of each analyte, respectively. The results obtained show that the sample volume (within the range studied) did not affect the preconcentration of the selected pesticides, except for metobromuron, for which a 20% decrease in preconcentration efficiency was observed when the sample volume was 50 ml. Selectivity: Application to Different Matrices The efficiency of the immunosorbent for the selective preconcentration of chlortoluron, isoproturon, metobromuron, linuron and chlorbromuron from ground, river and freeze-dried tap water samples was evaluated.Fig. 3 shows the chromatograms obtained from on-line preconcentration on the immunosorbent of 10 ml of (A) ground water and (B) river water spiked at the 0.05 mg l21 level with each phenylurea. It clearly demonstrates the high degree of clean-up obtained with the proposed method, since the baselines in the two chromatograms are comparable and as clean as that obtained by preconcentrating Milli-Q-purified water (see Fig. 1). This should allow calibration graphs to be constructed for all analytes in Milli-Q-purified water and to be used for any kind of water.Freeze-dried tap water sample containing pesticides with different chemical functionalities [atrazine and simazine (tria- Fig. 1 LC–UV trace obtained at 244 nm after on-line immunoextraction of 10 ml of Milli-Q-purified water spiked with 0.25 mg l21 of each phenylurea herbicide. Peaks: 1 = chlortoluron, 2 = isoproturon; 3 = metobromuron; 4 = linuron and 5 = chlorbromuron.The asterisk indicates an impurity arising from the synthesis of the immunosorbent. For LC conditions, see Experimental. Fig. 2 Binding curves for chlortoluron (1), isoproturon (2), metobromuron (3), linuron (4) and chlorbromuron (5) after on-line immunoextraction of 10 ml of Milli-Q-purified water sample when the compounds are alone (A) or mixed (B). See text for details. Analyst, October 1997, Vol. 122 1115zines), carbaryl (carbamate), propanil (propioanilide), linuron (phenylurea), fenamiphos (organophosphorus) and permethrin (pyrethroid)] at concentrations within the range 0.5–17.3 mg g21 was also preconcentrated in order to demonstrate the high selectivity of the immunosorbent.Fig. 4 shows the chromatograms obtained after on-line preconcentration of 10 ml of diluted reconstituted tap water sample (see Experimental), (A) on a precolumn packed with the apolar copolymer PRP-1 and (B) on the immunosorbent. The chromatogram is cleaner after preconcentration on the immunosorbent than on the PRP-1 precolumn, because the antibodies immobilised are only able to retain phenylurea herbicides.However, in Fig. 4(B), two peaks are observed, one at the retention time of linuron and another which was identified as propanil by constructing its specific calibration graph in Milli-Q-purified water using the procedure described. Although propanil belongs to another family of pesticides (propioanilides), there is a clear similarity between the structures of propanil and linuron, as shown in Fig. 4(B), and the antibodies are not able to differentiate between compounds with this high degree of structural similarity. This would make it possible to increase the number of compounds that the immunosorbent is able to retain, such as anilines, which are transformation products from the phenylurea herbicides and some of which are included in the list of priority pollutants to be monitored in environmental waters in Europe.This interesting possibility is now under study. Analytical Performance and Method Validation Calibration graphs for ground and river water were constructed by applying the on-line immunoextraction procedure to 10 ml of spiked water within the range 0.05–0.5 mg l21 and the resulting correlation coefficients were satisfactory (r2 > 0.99). The RSD for the selected analytes at a concentration of 0.25 mg l21 using the full procedure and evaluating the peak areas was in the range 1–9% (n = 5), depending on the pesticide.The detection limits, which were calculated as three times the standard deviation of the lowest concentration solutions, were within the range 0.01–0.03 mg l21, depending on the phenylurea herbicide, although they could be lowered further by preconcentrating a larger sample volume. These low detection limits obtained with only 10 ml of sample are a remarkable result and are good enough to allow the fate and transport of phenylureas to be studied directly in environmental waters.Calibration graphs were also constructed for propanil and linuron within the range 0.05–0.5 mg l21 by the on-line immunoextraction of 10 ml of Milli-Q-purified water and were used for the determination of propanil and linuron in the freezedried tap water samples. The results (Table 1) were in good agreement with the mean values obtained in an interlaboratory exercise,17 in which only validated methods were used.Reusability of the Immunosorbent The immunosorbent used in this study was employed in the analysis of more than 50 samples, including dirty water samples such as river water, and no decrease in the capacity of the immunosorbent was detected. This long lifetime is due to the good stability of the antibodies immobilised on aldehyde- Fig. 3 LC–UV traces obtained at 244 nm after on-line immunoextraction of 10 ml of ground (A) and river (B) water spiked with 0.05 mg l21 of each phenylurea herbicide.Peak numbers as in Fig. 1. For LC conditions, see Experimental. Fig. 4 LC–UV traces obtained at 244 nm after on-line preconcentration of 10 ml of freeze-dried reconstituted tap water sample on a PRP-1 precolumn (A) and on the immunosorbent (B). For LC conditions, see Experimental. Table 1 Mean values ± standard deviations (mg g21) obtained in the determination of propanil and linuron in freeze-dried tap water samples Interlaboratory On-line Compound exercise* immunoextraction† Propanil 11.70 ± 1.31 11.12 ± 0.24 Linuron 5.47 ± 0.90 5.66 ± 0.36 * Data obtained from ref. 17. † Mean value of three independent determinations. 1116 Analyst, October 1997, Vol. 122activated silica and because the regeneration (25 ml of PBS) and storage were appropriate for the period tested. Conclusion The use of a precolumn containing a mixed immunosorbent with immobilised antiisoproturon and antichlortoluron antibodies allows the simultaneous on-line immunopreconcentration of several phenylurea herbicides from ground and river water.The high selectivity shown by the immunosorbent allows the determination of phenylurea herbicides at trace levels when only 10 ml of water sample are percolated. Moreover, the selectivity of the immunosorbent was clearly demonstrated by analysing freeze-dried tap water samples spiked with large amounts of pesticides with different chemical functionalities, and the immunosorbent only retained linuron (phenylurea herbicide) and propanil (propioanilide), which show a high degree of structural similarity.Although this immunosorbent is not specific for isoproturon and chlortoluron, it offers the advantage of recognizing the phenylurea family of herbicides and very closely related compounds, which may be of great interest for environmental analysis. This work represents yet another example of the great potential shown by the immunosorbents used to date for the preconcentration of pesticides and further development in this area should be undertaken. This work received financial support from the Standards, Measurements and Testing Programme under contract number MAT1-CT940001 and PB95-0366-C02-01. The authors thank Max Gorman for revision of the manuscript. References 1 Barcel�o, D., and Hennion, M. C., Anal. Chim. Acta, 1995, 318, 1. 2 EC Directive relating quality of water intended for human consumption (80/778/EC), Off. J. Eur. Communities, L229/11, 1980. 3 van Ginkel, L. A., Stephany, R. W., van Rossum, H. J., and Zoontjes, P. W., Trends Anal. Chem., 1992, 11, 294. 4 Pichon, V., Chen, L., Hennion, M. C., Daniel, R., Martel, A., Le Goffic, F., Abian, J., and Barcel�o, D., Anal. Chem., 1995, 67, 2451. 5 Pichon, V., Chen, L., Durand, N., Le Goffic, F., and Hennion, M. C., J. Chromatogr. A, 1996, 725, 107. 6 Shahtaheri, S. J., Kwasowski, P., and Stevenson, D., J. Chromatogr. A, submitted for publication. 7 Shahtaheri, S. J., Katmeh, M .F., Kwasowski, P., and Stevenson, D., J. Chromatogr. A, 1995, 697, 131. 8 Thomas, D. H., Beck-Westermeyer, M., and Hage, D. S., Anal. Chem., 1994, 66, 3823. 9 Rollag, J. G., Beck-Westermeyer, M., and Hage, D. S., Anal. Chem., 1996, 68, 3631. 10 Lawrence, J. F., Menard, C., Hennion, M. C., Pichon, V., Le Goffic, F., and Durand, N., J. Chromatogr. A, 1996, 752, 147. 11 Lawrence, J. F., Menard, C., Hennion, M. C., Pichon, V., Le Goffic, F., and Durand, N., J. Chromatogr. A, 1996, 732, 277. 12 Rule, G. S., Mordehai, A. V., and Henion, J., Anal. Chem., 1994, 66, 230. 13 Mart�ýn-Esteban, A., Kwasowski, P., and Stevenson, D., Chromatographia, 1997, 45, 364. 14 Katmeh, M. F., Frost, G., Aherne, W., and Stevenson, D., Analyst, 1994, 119, 431. 15 Katmeh, M. F., Aherne, W., and Stevenson, D., Analyst, 1996, 121, 1699. 16 Katmeh, M. F., Thesis, Robens Institute, University of Surrey, Guidford, 1994. 17 Mart�ýn-Esteban, A., Fern�andez, P., C�amara, C., Kramer, G. N., and Maier, E. A., Int. J. Environ. Anal. Chem., in the press. 18 Pichon, V., Chen, L., and Hennion, M. C., Anal. Chim. Acta, 1995, 311, 429. 19 Price, C. P., and Newman, D. J., Principles and Practice of Immunoassay, Macmillan, London, 1991. Paper 7/02828H Received April 25, 1997 Accepted July 22, 1997 Analyst, October 1997, V

ISSN:0003-2654    

DOI:10.1039/a702828h

出版商:RSC    

年代:1997

数据来源: RSC