Recent developments in solid-phase microextraction coatings and related techniques

Journal of Chromatography A, 1103 (2006) 183–192

Review

Recent developments in solid-phase microextraction coatings and related techniques Christian Dietz, Jon Sanz, Carmen C´amara ∗ Department of Analytical Chemistry, Faculty of Chemistry, University Complutense de Madrid, Ciudad Universitaria, 28040 Madrid, Spain Received 2 August 2005; received in revised form 8 November 2005; accepted 14 November 2005 Available online 6 December 2005

Abstract During the last decade, solid-phase microextraction (SPME) has gained widespread acceptance for analyte matrix separation and preconcentration. Relatively few data are currently available dealing with in-house production of fibres with tailor-made properties to be used for SPME, though recently the number of publications evaluating new coatings has been considerably growing. This review, centred on publications that appeared during the last five years, is resuming different approaches which can be used for fibre production and further summarises alternative techniques closely related to SPME, such as in-tube extraction or single-drop microextraction (SDME). The aim is to give the reader a concise overview of recent developments in new coating procedures and materials, including the respective applications. © 2005 Elsevier B.V. All rights reserved. Keywords: Solid-phase microextraction; In-house fibre production; Fibre coating; SPME related techniques

Contents 1. 2.

3.

4.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Fibre coating procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. In-house fibre production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. Sol–gel technology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. In-tube extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Electrochemical procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Physical deposition. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. Direct use . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.7. On-fibre derivatisation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Alternative technique . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. Single-drop microextraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Liquid-phase microextraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Stir-bar sorptive extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.4. Thin-film microextraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

183 184 184 184 185 186 187 187 187 188 188 189 189 190 190 190

1. Introduction

∗

Corresponding author. Tel.: +34 91 394 4318; fax: +34 91 394 4329. E-mail address: [email protected] (C. C´amara).

0021-9673/$ – see front matter © 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.chroma.2005.11.041

Solid-phase microextraction (SPME) is a relatively recent technique [1], which was introduced in the early 1990s by Pawliszyn and Lord. It has been more and more widely used in sample preparation, especially since the first fibres became

184

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

commercially available in 1993 [2]. The most common applications, limited by the properties of commercially available fibres, are in the field of organic compounds, i.e. volatile organic compounds (VOCs), the group of benzene, toluene, ethylbenzene and xylenes (BTEX), polycyclic aromatic hydrocarbons (PAHs), pesticides, odour and flavour analysis, but recently even metallorganic compounds have been target analytes for this technique. Fields of application have been continuously growing, including environmental, clinical, forensic, food and botanical analysis, with nowadays about 3000 research papers published. Simultaneously the technique gained growing acceptance and increasing use in routine laboratories and industrial applications, especially since automatised fibre injection systems are on the market, normally hyphenated with gas chromatographic separation modules. The technique is based on the establishment of an equilibrium between the analyte and a fused silica fibre coated with a stationary phase, which can be a liquid polymer, a solid sorbent, or a combination of both. The analyte is then desorbed from the fibre into a suitable separation and detection system, usually a gas chromatograph. SPME increasingly substitutes classical and time consuming extraction and leaching processes. The main advantages of this technique are the simplicity of operation, its solventless nature, analyte/matrix separation and preconcentration. It covers a wide range of sampling techniques, including field, in situ and air sampling. Generally accepted drawbacks are a relatively poor reproducibility, lot-to-lot variations, lack of selectivity, sensibility against organic solvents, the latter frequently impeding liquid sampling and their cost. But possibly the most important disadvantage is the limited range of stationary phases which are commercially available, only roughly covering the scale of polarity. In particular these coatings are polydimethylsiloxane (PDMS), divinylbenzene (DB), polyacrylate (PA), Carboxen (CAR; a carbon molecular sieve) and Carbowax (CW; polyethylene glycol). Fibres are available in different coating combinations, blends or copolymers, film thickness, and fibre assemblies, enlarging to a certain extend the field of possible applications. The present review focuses on production processes and fibre characterisation in terms of polarity, active surface, porosity, film thickness, mechanical stability, etc.; rather than the on application to real samples and complete

method validation, thus the selected papers include sufficient details on fibre production to reproduce these coatings. 2. Fibre coating procedures 2.1. In-house fibre production To our knowledge there is only one research group dedicated to the entire production process of a fibre to be used for SPME purposes, which consists in a Nb(V) oxide coating [3], prepared by vapour deposition onto in-house made glass ceramic rods. The obtained fibres showed acceptable reproducibility and detection limits for determination of polar compounds such as alcohols and phenols [4], but were not yet applied to real samples. 2.2. Sol–gel technology The most common approach to overcome the mentioned limitations by tailor-made preparation of fibre coatings is to help oneself with sol–gel technology, which provides efficient incorporation of organic components into inorganic polymeric structures in solution under extraordinarily mild thermal conditions [5]. In general, the sol–gel process, involves the evolution of inorganic networks through the formation of a colloidal suspension (sol) and gelation of the sol to form a network in a continuous liquid phase (gel) [6,7]. Through this process, homogeneous inorganic oxide materials with desirable properties of hardness, chemical and thermal resistance, polarity and tailored porosity, can be produced at room temperatures. The precursors for synthesising these colloids consist of a metal or metalloid element surrounded by various reactive ligands. Metal alkoxides, such as the alkoxysilanes tetramethoxysilane (TMOS) or tetraethoxysilane (TEOS) are most popular because they react readily with water. To describe the sol–gel process at the level of functional groups, three consecutive reactions, as shown in Fig. 1, are generally used: hydrolysis (R1 in Fig. 1), alcohol condensation (R3 in Fig. 1) and water condensation (R2 in Fig. 1). Generally speaking, reaction R1 replaces alkoxide groups with hydroxyl through the addition of water. Subsequent condensation reactions (R2 and R3) involving the silanol

Fig. 1. Basic steps in sol–gel coating technology.

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

groups produce siloxane bonds plus the by-products water or alcohol. Under most conditions, condensation commences before hydrolysis is complete, though choice of the catalysator may inverse this observation. Additionally, because water and alkoxides are immiscible, a mutual solvent such as an alcohol as homogenizing agent. Here hydrolysis is facilitated due to the miscibility of the alkoxide and water. As the number of siloxane bonds increases, the individual molecules are bridged and jointly aggregate in the sol, which is a colloid suspension of solid particles of a diameter of few hundred of nanometers in a liquid phase. When the sol particles aggregate, or cross-link into a network, a macromolecular gel is formed. Upon drying, trapped volatiles are then driven off. Thin films can be produced on a piece of substrate by dip-coating, which is the most popular strategy for SPME fibre production. Polyethyleneglycol coatings have been basically used for BTEX determination [8–10]. PDMS as coating has been applied to dextromethorphan [11] and selenoaminoacid determination [12], the latter after derivatisation with isobutylchloroformate. Mixed polymers of PDMS and polyvinyl alcohol could be proven to be useful in determination of pesticides [13], PAHs [14] and related reagents. Phenyltrimethoxysilane [15] has been attached to an optical fibre support and applied to non polar compounds (benzene, toluene, ethylbenzene, 2-octanone, benzaldehyde, acetophenone, dimethylphenol and tridecane). Another group of coatings belong to the not commercially available stationary phases, for example a variety of crown ether coatings, which have been used for determination of BTEX [16,17], organophosphorous pesticides [18], aromatic amines [19] and phenolic compounds [20]. Different calyx[4]arene compounds were applied to the determination of chloro-containing pesticides [21], chlorophenols [22], phenolic compounds [23] and PAHs [24]. Carbon based material such as glassy carbon were found to be suitable for BTEX [25] and monohalogenated benzenes [26], for the latter low temperature fluorinated glassy carbon was synthesised in order to achieve selective extraction. Hydroxyfullerene coatings [27] were applied to polychlorinated biphenyls (PCBs) and PAHs. Less currently used sol–gel based coatings are 3(trimethoxysilyl)propyl methacrylate [28], which been used for determination of aroma compounds in beer. Three more types of acrylate/silicone co-polymers have been prepared and characterised using the sol–gel technology and cross-linking method, these coatings have been found to be solvent resistant, thermally stable up to 350 ◦ C and showed lifetimes higher than those of commercial fibres [29]. One those, in particular co-poly(butyl methacrylate/hydroxy terminated silicone oil was found to be optimum for the determination of 2-chloroethyl ethyl sulphide, a surrogate of the chemical warfare agent mustard gas, in soil samples. Another “exotic” coating is C8 -TEOS [30], which has been, in combination with HPLC, applied for determination of phenylated species of Arsenic, mercury and tin. A very interesting study about synthesis, characterisation and properties of new polyfunctional sol–gel precursors has recently appeared [31], though the targeted applications in this case were more focused on enhancement of abrasive and chemical resistance of the obtained coatings. However, the characteristics and properties

185

of a particular sol–gel coating are strongly related to a number of factors, such as, H2 O/Si molar ratio (R), pH, nature and concentration of catalyst, temperature and time of reaction, ageing temperature and time and drying during the synthesis of the coating. The lack of standardised procedures in this field causes problems in inter laboratory reproducibility. 2.3. In-tube extraction The frontier in between sol–gel chemistry and alternative methods can be drawn in the field of in-tube extraction (ITE). This technique can be defined by the use of an open tubular fused-silica capillary column as an extraction device, analytes in aqueous samples are directly extracted and concentrated into the stationary phase of capillary columns by repeated draw/eject cycles of sample solution, which in turn can be directly transferred to a liquid chromatographic column. The stationary phase can be either prepared by sol–gel technology, alternative chemical coating procedures or selected out of the range of commercially available GC capillary columns. Examples for the first case are the synthesis of a zirconia-based hybrid organic–inorganic sol–gel coating [32], which has been applied to the determination of PAHs in combination with GC, or a ␤-cyclodextrin coating [33], used for the determination of non-steroidal antiinflammatory drugs. Another hybrid material to be used as sorbent in ITE is TiO2 -PDMS [34], a capillary coated with this technique was applied to the determination of PAHs, ketones, and alkylbenzene in aqueous samples. An interesting approach is the use of functionalised mesoporous silica for ITE purposes [35], where the inner surface of the capillary was first coated with the ordered mesoporous silica film by sol–gel method and then chemically modified with octadecyltrimethoxysilane. Improved extraction efficiencies for biphenyl were reported when the mesostructural coating was more ordered. Commercial GC columns such as Omegawax [36,37] have been used in combination with HPLC detection for determination of benzene and ␤-blockers in urine, respectively. A porous divinylbenzene polymer containing column was applied to the determination of five estrogens in river water [38], here the extraction step was on-line coupled to LC/MS/MS detection. PDMS columns packed with either Zylon (a fibrous rigid-rod heterocyclic polymer; poly(p-phenylene-2,6-benzobisoxazole) fibres [39] or stainless steel wire [40], the latter in order to reduce the internal volume, have been applied to the determination of antidepressant drugs. Although the length of the capillary in ITE is normally short, up to 1 m columns [41] (0.25 i.d., f = 3.5 ␮m PDMS) have been used, quantitative back extraction of some BTEX compounds could be achieved with this approach. Amphiphilic and hydrophilic oligomers were attached by the sol–gel method to fused-silica capillaries [42] and applied to the determination of a variety of organochlorine pesticides, triazine herbicides, estrogens, alkylphenols and bisphenol-A, which can be considered as an example of a mixed application of SPME and ITE, as extraction was carried out in headspace. As the volume of extraction sorbent in ITE is generally small, much effort was spent in developing sorbents with enhanced extraction efficiency or selectivity. One approach here is the use of

186

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

molecular imprinted polymers (MIPs), in particular methacrylic acid as monomer and ethylene glycol dimethacrylate as crosslinker, which was, packed in polyether ether ketone (PEEK) columns [43] and used for determination of ␤-blockers from biological fluids. The same column, packed with alkyl-diol-silica particles [44], simultaneously showed good anti-fouling properties and high extraction efficiency for benzodiazepines, which makes the method suitable for direct use with serum samples, without need for ultrafiltration or other deproteinization steps prior to handling the biological sample. Zylon has been used in the same way for determination of butylphthalate in wastewater [45] and including for on-line coupling of ITE and capillary electrophoresis [46] for determination of tricyclic antidepressants. Another way to increase the extraction efficiency is the use of polymer monolithic material, whose double-pore structure offers convective mass transfer, preferable over diffusive mass transfer in extraction processes. Both the through-pores and the mesopores structures can be optimised for ITE purposes. This material can be easily in situ synthesised, initiated thermally or by radiation, using monomer mixture, cross-linker and proper porogenic solvent in a “mold”. It generally provides biocompatibility and pH-stability, which makes these columns suitable for use with biological samples such as urine or serum. A C18 bonded monolithic silica column, prepared by in situ hydrolysis and polycondensation of alkoxysilane [47], showed preconcentration efficiencies higher than those obtained by conventional ITE for pesticides [48] and a range of other common analytes such as toluene, naphthalene, biphenyl and fluorene. A poly(methacrylic acid-ethylene glycol dimethacrylate) monolithic capillary column has been prepared and evaluated for ketamine [49], theobromine, theophylline and caffeine in human serum [50] and amphetamine in urine samples [51]. Chemically or electrochemically deposited polypyrrole (PPY) coatings have been coupled to either HPLC [52] or IC [53] for determination of aromatics and anions, respectively. Sensitivity and selectivity of these coatings can be tuned for both, in-tube extraction and SPME fibres, by simple selection of the film thickness [54]. A review on details for in-tube extraction

technique and its most common applications has been given by Kataoka [55]. 2.4. Electrochemical procedures The next group of alternatives for fibre production is based on electrochemical methods for deposition of a coating on the support material, which in this case has to be conductive. The most popular approaches are potentiometry and cyclic voltamperometry (CV), in the latter the potential is changed gradually and repeated like a sawtooth waveform as the potential measured between the reference electrode and the working electrode and the current measured between the working electrode and the counter electrode are plotted. The potential is changed linearly and the scan rate can be changed, so compounds are oxidised and reduced reversibly on the electrodes. A simple electrochemical cell to be used for this purpose consists of three electrodes, due to the difficulties arising of the concurrent measurement of current and potential. The first those is the working electrode, which is the electrode at which the electrochemical phenomena (reduction or oxidation) being investigated are taking place. The second is the reference electrode, whose potential is constant enough that it can be taken as the reference standard against which the potentials of the other electrodes present in the cell can be measured. Commonly used reference electrodes are the silver–silver chloride electrode (Ag/AgCl/4 M KCl, e = 0.222 V) or the Calomel electrode (Hg/HgCl/KCl). The third electrode is the auxiliary electrode, which serves as a source or sink for electrons so that current can be passed from the external circuit through the cell. In Fig. 2a, the practical set-up of a such threeelectrode system is shown, Fig. 2b shows the correspondent voltammogram for a typical electrocoating process applying five oxidation/reduction cycles. Platinum wire has been used as support material for preparation of dodecylsulfate-doped polypyrrole [56] and poly3-methylthiophene [57] films. Like in the latter case, where arsenate has been determined, the change of a potential is frequently used as driving force for an analyte extraction/desorption

Fig. 2. Three-electrode configuration used during an electrocoating process (A) and corresponding cyclic voltammogram (B) for five cycles applied (arrow indicates direction of the scans).

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

process. Poly-3-dodecylthiophene [58] has been used in a similar way for preconcentration of arsenobetaine from aqueous samples, preconcentration of this neutral species was achieved in normal SPME immersed mode and desorption by applying an external potential. Another example for this technique is the determination of nickel and cadmium ions on an overoxidised sulfonated polypyrrole (OSPPy) film [59,60]. Tailored properties of polypyrrole coatings can be achieved by proper selection of the counterion, this dependence has been studied for the determination of VOC’s in headspace mode and for polar and ionic compounds in aqueous solution [61]. Electroplating of a 10 ␮m gold coating on a carbon steel wire enabled mercury determination [62] by electrodeposition at −0.2 V and subsequent capacitive discharge desorption onto a GC–MS. On the other hand, electrochemically prepared fibres can as well be used in the normal SPME headspace extraction mode. Cyclic voltammetry has been used to deposit polyaniline (PANI) on platinum wire [63] for the determination of phenolic compounds in water. This polymer exists in a variety of protonation and oxidation forms, in particular leucoemeraldine (fully reduced form), emeraldine and pernigraniline (fully oxidised form). These forms can be specifically achieved by adjusting the scan rate and swept potential [64] during the electrocoating process using CA. For the determination of anatoxin-a, an alkaloid, produced by cyanobacteria, from cultured media and contaminated water by SPME-GC–MS, the leucoemeraldine base form of PANI as fibre coating revealed highest sensitivity. The same coating can be as well achieved by the application of a constant voltage, which has been done for a steel [65] and gold [66] support, these fibres have been used for the determination of six aromatic amines and aliphatic alcohols from gaseous samples, respectively. An interesting approach is potentiometrically deposited Cu(I) on a copper wire, which has been applied to amine determination [67], here the extraction mechanism seems to be due to adsorption only. 2.5. Physical deposition Subsequent dipping of the support material into polymer solution, frequently followed by some kind of thermal treatment or conditioning procedure for stabilisation of the polymer, is a very fast and comfortable way of obtaining fibre coatings, though the resulting fibres mostly suffer from mechanical instability. This factor can be that important that the film has to be classified as “disposable” and to be renewed prior to each analysis, which is the case of an ionic liquid coating [68] applied to BTEX analysis. Other polymer films prepared in this way are based on Nafion [69], used for determination of polar solvents in liquid matrix, and MIPs [70], the latter applied to the determination of brombuterol. A mechanically very stable SPME fibre could be achieved by introducing a silver wire into a suspension of alumina and polyvinyl chloride (PVC) powder as matrix dispersed in tetrahydrofuran (THF) [71], the porous coating was stable up to 300 ◦ C and applied to alcohol extraction from fruit juices and vinegar. As an “exotic” application of this method can be considered the use of an ion exchanger, di(2-ethylhexyl)phosphoric acid [72], applied to Bi(III) extraction followed by desorption into KI and detection by UV–vis.

187

Attaching a porous material onto a support by using glue (epoxy or similar) is another possibility to obtain fibres for SPME applications. In that way mesoporous silica particles attached to a stainless steel fibre could be used for determination of aromatic hydrocarbons [73,74]. A problem with this coating arises due to that mass transfer from bulk solution to the small mesopores is diffusion controlled, thus stirring rate and temperature during sampling strongly influence the extraction efficiency. Nevertheless, the approach seems to be promising as adsorption capacity is pore size controlled and absorption can be achieved by tailor-made functional groups incorporated during synthesis of the particles. The same support was covered with silicon particles bounded with phenyl-C18 and C15 material, normally used as HPLC stationary phases, for extraction of BCPs from water and PAHs from soil [75]. Another group reported to have used activated charcoal onto an optical fibre [76] for BTEX determination, but no details on the coating process are given. A stainless steel wire, immersed into a slurry formed of an ion exchange diol silica material (XDS), epoxy and chloroform and then baked at 180 ◦ C [77], resulted in a SPME fibre capable to selectively extract small peptides from matrices as complex as whole blood. The fibre was stable for about 150 extraction/desorption cycles with a reproducibility of 5.6%, with enough sensitivity to work at physiological levels for determination of angiotensin 1 and angiotensin 2. 2.6. Direct use Some materials even seem to have properties which allow their direct use for some, though punctual applications. Polyimide, which is the standard coating for optical fibres in order to enhance mechanical stability, has been used for Cr(III) extraction after derivatisation with TFA [78] and for chlorinated hydrocarbons as well as PCBs [79], in the latter case the behaviour of an uncoated fused-silica fibre has as well been evaluated. HBtype pencil lead has been applied to the determination of three pesticides [80], but the material showed severe carryover effects. Another graphite material has been applied to determination of surfactants [81], though in this case it cannot be considered as a real SPME application because the fibre served as probe for direct electrospray formation. Anodised aluminium wire was found to be useful for extraction of aliphatic alcohols and BTEX from gaseous samples [82]. 2.7. On-fibre derivatisation In order to improve separation, selectivity or sensitivity and to extend the range of possible target analytes in SPME, frequently analyte derivatisation is required. This can be done in solution prior to sampling, during desorption in the GC injection port or directly on the solid phase of the extraction fibre, avoiding possible side reactions and interferences. The technique will here only be briefly treated, due to that mostly commercial fibre coatings are employed. Early work in this field was carried out by Pawliszyn’s group, with a detailed study of different approaches for derivatisation targeting fatty

188

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

acids [83]. For on-fibre derivatisation, the derivatising agent has to be loaded on the fibre coating prior to analyte sampling, which is normally done by exposing the coating to vapour of the respective reagent, an alternative approach is to immerse the fibre into a volatile solvent in which the reagent has been dissolved. Using the first method trifluoroacetic anhydride (TFA) was used for derivatisation of amphetamines [84], previously loaded onto a PDMS fibre. The same analyte class and methamphetamine in urine were determined using a 4:1 mixture of heptafluorobutyric anhydride and heptafluorobutyric chloride as derivatising reagents [85], here the PDMS fibre was directly exposed to the headspace of the analyte reagent mixture. A selection of commercial fibres were proven to be useful for determination of methylamine when on-fibre derivatised with 9-fluorenylmethylchloroformate [86]. A PA-coated fibre showed to be suitable for determination of acidic herbicides [87] after derivatisation with diazomethane. The same fibre provided enhanced extraction efficiency for estrogens when methyl-N(trimethylsilyl)trifluoroacetamide [88] as derivatisation reagent was present in the headspace. A related silylation agent, Nmethyl-N-(tert-butyldimethylsilyl)trifluoroacetamide [89], was used for acidic herbicides, the PA fibre was found to be optimum to carry out the derivatisation and subsequent trapping of the reaction products. Bis(trimethylsilyl)trifluoroacetamide [90] was applied for silylation of hydroxy metabolites of PCBs in urine, here the fibre of choice to carry out the reaction was PDMS/DVB. For carbonyl-containing compounds the most popular approach is the use of oxime forming o-(2,3,4,5,6pentafluorobenzyl)-hydroxylamine (PFBHA) as derivatising agent in combination with a PDMS/DVB fibre, in this way formaldehyde in air [91], atmospheric metabolites of volatile compounds exhaled by plants [92], aldehydes in water [93] and beer [94], hexanal and heptanal in human blood [95] and acetone in human breath [96] were determined. A commercial automated system using a method based on the same derivatisation reagent for determination of C1 –C10 aldehydes [97] in particleboard, wine and fish samples, was recently evaluated. Limits of detection (LODs) achieved for these compounds were at the sub ␮g L−1 level. Further hydrazon forming pentafluorophenylhydrazine can be used with this group of analytes, which has been done to determine the carbonyls generated during the thermal degradation of sunflower oil [98]. These methods are considered to be inferior to competing techniques, such as solid-phase extraction, due to the small amount of analyte which can be extracted and additional instrumentation required. A review on derivatisation techniques for SPME has been recently given by Stashenko and Martinez [99]. 3. Alternative technique 3.1. Single-drop microextraction Recently, alternative but SPME related concepts have been introduced, which will be also addressed. Single-drop microextraction (SDME) [100,101] involves the extraction of organic

contaminants in headspace or aqueous sample solution into a microdrop of a water immiscible organic acceptor solvent, commonly a few microliters, which is suspended on the tip of a microsyringe. Unlike fibres, drops can be renewed for each extraction and it is an elegant method to overcome the limited availability of fibre coatings, as a wide variety of solvents and trapping agents can be used. A review on SDME, including 27 references [102], resumes the investigation carried out in this nowadays fast growing field until 2002. In more recent work, toluene has been used for extraction of organo-P insecticides [103], nitroaromatic explosives [104], and chlorobenzenes [105], respectively. A 3:1 mixture of chloroform/toluene was employed for extraction of 12 amino acids [106], after their derivatisation into the correspondent N(O,S)-ethoxycarbonyl amino acid ethyl esters. Isooctane as solvent has been employed for endosulfan [107] and iodine determination [108], the latter after derivatisation. Hexane was found suitable for extraction of antifouling agents [109], organophosphorous insecticides [110], mixed with dichloromethane it was used for determination of dialkyl phthalate esters [111]. Cyclohexane was employed for extraction of essential oil components such as camphor, borneol [112] or panaxynol [113] from traditional Chinese medicine after previous pressurised hot water extraction. A benzyl alcohol microdrop was found to be the optimum solvent for extraction of 2-butoxyethanol [114], an emulsifier used in colour industry. Octanol provided optimum extraction efficiency for a variety of short chain alcohols [115] from water. The field of applications has been even extended to the determination of metal-organic compounds, such as tributyltin [116], which was extracted into a decane microdrop. Methylmercury [117] was trapped as hydride into a microdrop containing finely dispersed Pd(0) and Pt(0), species which formed in situ from metal salts under the conditions of hydride generation inside the drop. A Pd2+ containing microdrop, acidified with HNO3 , showed to be highly effective for trapping of other hydride forming elements [118] such as arsenic, selenium, and antimony, achieving sub-ppb limits of detection and relative standard deviations around 3%. Introducing a microliter-size liquid membrane in between sample and microdrop, the former consist of octane and the latter of water, a simultaneous extraction/back extraction process was reported [119], achieving enrichment factors of up to 500 for some amphetamines. Higher concentration factors than in the static mode, where extraction is generally carried out in a septum closed vial, could be achieved in a continuous mode, where the sample solution flows around the suspended microdrop. Using hexane as solvent, volatile halohydrocarbons could be extracted in this way from aqueous samples [120], with LODs as low as 0.001 ␮g L−1 for CCl4 , using GC with electron-capture detection. An ionic liquid (1octyl-3-methylimiazolium hexafluorophosphate) as extraction solvent was found to be suitable for extraction of substituted phenols [121] and for formaldehyde in shiitake mushrooms [122], after derivatisation with 2,4-dinitrophenylhydrazine. In spite of being a virtually solventless, inexpensive, fast and simple method for analyte extraction and/or preconcentration, frequently problems with drop stability and lack of sensitivity have been reported, a further development trying to overcome

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

these limitations is hollow fibre liquid-phase microextraction (LPME). 3.2. Liquid-phase microextraction The use of a porous polypropylene hollow fibre connected from one end to the tip of a microsyringe added a protective feature to the organic phase and served as an interface between the donor and acceptor phases. Polypropylene is commonly used because it is highly compatible with a broad range of organic solvents. This technique [123], which can be considered as a further development of SDME, is known as LPME. Illustratively the organic phase is sandwiched between two aqueous phases, one outside the fibre and the other phase is inside the lumen of the fibre. The analytes partition from the aqueous donor phase into the organic phase and subsequently partition into the aqueous acceptor phase where they are concentrated. The possibility of accommodating larger volumes of acceptor solvent phase results, in general, in increased sensitivity and reduces carry-over effects. Toluene is one of the most commonly used acceptor phases, which ideally should provide good immobilisation in the hollow fibre pores, high solubility for the target analyte, be immiscible in water and stable enough over the extraction time. Based on this solvent as acceptor phase, two methods were validated and applied to the determination of low molecular weight PAHs [124,125] and organophosphorous insecticides, respectively, in water samples. Eight other pesticides were extracted from aqueous soil samples [126] without any further sample preparation. The same solvent was used for enantiomeric extraction of the antidepressant mirtazapine [127] from human plasma, followed by separation on a chiral column. Twenty persistent organic pollutants [128] such as organochlorine pesticides and PCBs could be extracted from marine sediments after microwave assisted extraction into water. Vinclozolin [129], a dicarboximade fungicide which may act as hormone mimic or antagonist, was equally preconcentrated from sediment and water samples. Six more fungicides [130] could be equally determined in the same sample matrix, obtaining enrichment factors ranging from 139 for procymidone to 213 for chlorothalonil. Another application of toluene in LPME is the extraction of alkylphenols, chlorophenols and bisphenol-A in aqueous samples [131], here silylation in the injection port of the GC is required prior to analyte separation. Direct coupling to GC was achieved for the determination of nitroaromatic degradation products of explosives in water [132] and for determination of PCBs [133] in human serum. Octanol was employed for extraction of acidic pharmaceuticals (ibuprofen, diclofenac, etc.) from wastewater [134], for methamphetamine, preconcentration factors of 75 were achieved using a fibre loaded with this solvent and an acidic acceptor solution [135]. This acceptor phase was further found to be suitable for preconcentration of hydroxyl metabolites of PAHs [136] and for determination of the mycotoxin ochratoxin A in wine [137]. For weakly basic drugs (benzodiazepines and non-benzodiazepine) [138], a study revealed decreasing extraction efficiency the longer the alkyl chain of the alcohol used as organic phase was. Nevertheless, nonanol was chosen as optimum, as with octanol

189

and extraction from whole blood matrix, the system was found to be not robust enough. Adding crown ethers as complexing agents for aromatic amines [139] to the acceptor phase resulted in accelerated extraction times. Carrier-mediated LPME, where analytes are extracted as ionpairs through a thin layer of a water immiscible organic solvent immobilised in the pores of the porous hollow fibre (liquid membrane), and the into small volume of an acidic aqueous acceptor solution placed inside the lumen of the hollow fibre [140], was found to be suitable for extraction of polar drugs from human plasma. Recently, a two-step LPME method was introduced, achieving enrichment factors above 6000 for aromatic amines [141]. The first step involved a benzyl alcohol/ethyl acetate (80/20%, v/v) loaded polypropylene membrane as interface between aqueous donor and acceptor phase, the acceptor solution in the first step was the donor phase for the second step, employing a polypropylene hollow fibre, as acceptor phase. Instead of the latter material, packed columns can also be used in LPME, e.g. polytetrafluoroethylene fibres loaded with hexane [142] showed acceptable extraction behaviour for BTEX and similar compounds. A slightly different approach is to coat this kind of hollow fibre material and to use it directly for extraction. This has been done employing amphiphilic polyhydroxylated polyparaphenylene [143] as coating for determination of organochlorine pesticides in water. Desorption was carried out in hexane, but the method seems to be limited because the polymer is soluble in tetrahydrofuran, chloroform, toluene, dichloromethane and dimethylformamide. The disposable nature of the hollow fibre eliminates the possibility of carry-over effects and cross-contamination, thus providing enhanced reproducibility. Further, the small pore size prevents large molecules and particles present in the donor solution from entering the acceptor phase, providing effective matrix/analyte separation. Details on hollow fibre configurations, LPME sampling modes and different parameters to be taken into account during method optimisation have been reviewed in 2003 by Psillakis and Kalogerakis [144], including 45 references. Another review, including 33 references and appeared in 2004, focuses on the use of LPME for drug analysis [145]. A insight in basic extraction principles, technical set-up, recovery, enrichment, extraction speed, selectivity and applications can be found in a review given by Rasmussen and Pedersen-Bjergaard [146], including 41 references. A frequently reported drawback of this technique is a lack of precision, which may be attributed to the completely manual operation from fibre preparation and conditioning to the handling of small extract volumes. 3.3. Stir-bar sorptive extraction The amount of analyte extracted in SPME is proportional to the volume of the extraction phase, hence the sensitivity of a method can be improved by increasing the volume of the extraction phase. This can be achieved by an increase of the film thickness, either on a common fibre support, by coating onto thicker supports or by using the multifibres method. However, much longer equilibration time is required because the

190

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

extraction rate is controlled by the diffusion from the sample matrix through the boundary layer to the extraction phase. In general, SBSE is considered to be superior to SPME in terms of sensitivity and accuracy for determinations at trace level in difficult matrices, for example organophosphorous pesticides in honey [147]. Here, like in most other applications, a commercially available 500 ␮m–1 mm PDMS-coated stir bars was used. A review on SBSE for volatile and semivolatile components [148] of biological matrices was recently given. In the following only in-house developed procedures for stir-bar coating will be taken into consideration. Details on parameters affecting the production process of a partially cross-linked PDMS coating on glass stir bars by sol–gel technology were evaluated [149], using PAHs, n-alkanes and organophosphorous pesticides a model analytes. Temperature of the ageing step was found to be of special importance to avoid cracking of the film, finally a compact and thermally stable porous hydroxy-terminated sol–gel network with 30 ␮m film thickness was achieved in one coating process. A simple PDMS rubber tubing mounted onto a glass rod [150] was equally found to be suitable for automated sampling/desorption of a standard mixture of 44 organic compounds, including PAHs, phthalates, substituted benzenes and other. A similar approach, but including chemical bonding of a 500 ␮m PDMS film onto a titanium tube [151], was used for herbicide extraction. A normal polytetrafluoroethylene-coated stir bar [152] was evaluated for sorptive extraction using phenanthrene as model analyte, extraction was carried out into acetonitrile using ultrasonic energy. No memory effects were observed but the equilibrium times which had to be applied were rather high and standard deviation under these conditions well above 10%. An alkyldiol-silica (ADS) restricted access material, glued with epoxy resin on a magnet containing sealed glass tube [153] was used for extraction of caffeine and metabolites from biological fluids. In spite of surface investigations using SEM, the film thickness was not determined, the coating was found to be suitable for extraction of the target analytes from plasma without need for prior protein precipitation. One drawback of SBSE is the desorption step, as the analyte loaded on the coated stir bars cannot be desorbed directly in the injection port of a GC equipment. Hence the analyte has to be back extracted into a fitting solvent, which adds an additional step to the overall analytical method, or a specially designed thermal desorption unit has to be used. This desorption unit is in general relatively sophisticated instrumentation due to problems with high dead volume, further, the stir bar has to be manually transferred to the desorption unit. This may cause partly loss of the sensitivity gained by the use of extended sorbent surface. 3.4. Thin-film microextraction Another approach to obtain a higher volume of the extraction phase is extending the surface area of the polymer, which has been done by using membranes instead of fibre coatings. The use of a thin membrane has the advantage that enhanced extraction efficiency and hence high sensitivity can be achieved without suffering from elevated equilibrium times, as happens using thick phase coated stir bars. A cross-linked commercial

PDMS membrane [154] was successfully evaluated for extraction of PAHs in headspace mode. An in-house prepared membrane of PDMS/␤-cyclodextrin [155] showed good solvent resistance, uniform structure and was applied to both non polar PAHs extraction and to polar phenolic compounds. A commercial porous polysulfone hollow fibre membrane was coated with a variety of hydroxylated polymethacrylate compounds [156], resulting in a coating with an elevated number of functional groups, resulting in a membrane with increased swelling tendency when used in water. This proved to be an advantage over classical SPME for extraction of alkyl substituted phenols in seawater samples. A general drawback here is the lack of high volume GC injection or thermal desorption systems, thus for desorption the membrane usually is removed from the sample solution by tweezers, air dried, cut into pieces and the transferred to a vial containing a suitable solvent. 4. Conclusions In contrary to commercially available fibres for SPME, where for a limited number of coatings an extensive number of applications are reported, there are a high number of home-made coatings to be found in the literature, in their vast majority relatively unexploited from the application side of view. Additionally, at least four SPME-related techniques, such as LPME or SBSE, have been recently introduced to the analytical toolkit. Doubtlessly, a lot of in-house made fibres still suffer from problems like lack of mechanical or thermal stability, but the general impression is that there exists a wide range of possible solutions for a given analytical problem in SPME. Much of the techniques presented can be considered as complementary, e.g. research using SDME could easily establish the potential of a new, liquid fibre coating for a certain target analyte. In case of promising results here one could select out of at least seven completely different approaches for subsequent, tailor-made fibre production to be used for further method optimisation and validation. High surface sorptive techniques such as SBSE or the use of membranes may be an alternative to SPME for very low analyte concentrations, though high volume desorption is somewhat troublesome. Last but not least, most described fibres or correspondent extraction devices can be produced at a cost inferior to 5 D /unit in less than 15 min. References [1] H. Lord, J. Pawliszyn, J. Chromatogr. A 885 (2000) 153. [2] J. Pawliszyn, US Patent 5 691 206 (1997). [3] C. Berto da Silveira, A. Fernandes de Oliveira, S. Denofre de Campos, ´ Antˆonio de Campos, E. Carasek, Colloid Surf. A 259 (2005) 15. E. [4] A. Fernandes de Oliveira, C. Berto da Silveira, S. Denofre de Campos, ´ Antˆonio de Campos, E. Carasek, Talanta 66 (2005) 74. E. [5] S.L. Chong, D. Wang, J.D. Hayes, B.W. Wilhite, A. Malik, Anal. Chem. 69 (1997) 3889. [6] R. Zusman, I. Zusman, J. Biochem. Biophys. Methods 49 (2001) 175. [7] H. Wada, K. Kotera, H. Matsuura, K. Jinno, Y. Saito, Patent EP1411068-A, 2004. [8] Z. Wang, C. Xiao, C. Wu, H. Han, J. Chromatogr. A 893 (2000) 157. [9] Z. Wang, C. Xiao, C. Wu, H. Han, J. Chromatogr. A 893 (2000) 157. [10] R.G. da Costa Silva, F. Augusto, J. Chromatogr. A 1072 (2005) 7.

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192 [11] H. Bagheri, A. Es-haghi, M.-R. Rouini, J. Chromatogr. B 818 (2005) 147. [12] A.P. Vonderheide, M. Montes-Bayon, J.A. Caruso, Analyst 127 (2002) 49. [13] V.G. Zuin, A.L. Lopes, J.H. Yariwake, F. Augusto, J. Chromatogr. A 1056 (2004) 21. [14] A.L. Lopes, F. Augusto, J. Chromatogr. A 1056 (2004) 13. [15] M. Azenha, C. Malheiro, A.F. Silva, J. Chromatogr. A 1069 (2005) 163. [16] D. Wang, J. Xing, J. Peng, C. Wu, J. Chromatogr. A 1005 (2003) 1. [17] L. Yun, Anal. Chim. Acta 486 (2003) 63. [18] J. Yu, C. Wu, J. Xing, J. Chromatogr. A 1036 (2004) 101. [19] Z. Zeng, W. Qiu, M. Yang, X. Wei, Z. Huang, F. Li, J. Chromatogr. A 934 (2001) 51. [20] Z. Zeng, W. Qiu, Z. Huang, Anal. Chem. 73 (2001) 2429. [21] C. Dong, Z. Zeng, X. Li, Talanta 66 (2005) 721. [22] X. Li, Z. Zeng, J. Zhou, Anal. Chim. Acta 509 (2004) 27. [23] F. Zhou, X. Li, Z.O. Zeng, Anal. Chim. Acta 538 (2005) 63. [24] X. Li, Z. Zeng, S. Gao, H. Li, J. Chromatogr. A 1023 (2004) 15. [25] M. Giardina, S.V. Olesik, Anal. Chem. 73 (2001) 5841. [26] M. Giardina, L. Ding, S.V. Olesik, J. Chromatogr. A 1060 (2004) 215. [27] J. Yu, L. Dong, C. Wu, L. Wu, J. Xing, J. Chromatogr. A 978 (2002) 37. [28] M. Liu, Z. Zeng, B. Xiong, J. Chromatogr. A 1065 (2005) 287. [29] M. Liu, Z. Zeng, H. Fang, J. Chromatogr. A 1076 (2005) 16. [30] T.P. Gbatu, K.L. Sutton, J.A. Caruso, Anal. Chim. Acta 402 (1999) 67. [31] S. Hofacker, M. Mechtel, M. Mager, H. Kraus, Prog. Org. Coat. 45 (2002) 159. [32] K. Alhooshani, T.-Y. Kim, A. Kabir, A. Malik, J. Chromatogr. A 1062 (2005) 1. [33] Y. Fan, Y.-Q. Feng, S.-L. Da, Z.-H. Wang, Talanta 65 (2005) 111. [34] T.-Y. Kim, K. Alhooshani, A. Kabir, D.P. Fries, A. Malik, J. Chromatogr. A 1047 (2004) 165. [35] Y. Fan, Y.-Q. Feng, Z.-G. Shi, J.-B. Wang, Anal. Chim. Acta 543 (2005) 1. [36] R. Eisert, J. Pawliszyn, Anal. Chem. 69 (1997) 3140. [37] H. Kataoka, S. Narimatsu, H.L. Lord, J. Pawliszyn, Anal. Chem. 71 (1999) 4237. [38] K. Mitani, M. Fujioka, H. Kataoka, J. Chromatogr. A 1081 (2005) 218. [39] M. Imaizumi, Y. Saito, M. Hayashida, T. Takeichi, H. Wada, K. Jinno, J. Pharm. Biomed. Anal. 30 (2003) 1801. [40] Y. Saito, M. Kawazoe, M. Hayashida, K. Jinno, Analyst 125 (2000) 807. [41] B.C.D. Tan, P.J. Marriott, H.K. Lee, P.D. Morrison, Analyst 124 (1999) 651. [42] C. Basheer, S. Jegadesan, S. Valiyaveettil, H. Kee Lee, J. Chromatogr. A 1087 (2005) 252. [43] W.M. Mullett, P. Martin, J. Pawliszyn, Anal. Chem. 73 (2001) 2383. [44] W.M. Mullett, K. Levsen, D. Lubda, J. Pawliszyn, J. Chromatogr. A 963 (2002) 325. [45] Y. Saito, Y. Nakao, M. Imaizumi, T. Takeichi, Y. Kiso, K. Jinno, Fresenius J. Anal. Chem. 368 (2000) 641. [46] K. Jinno, M. Kawazoe, Y. Saito, T. Takeichi, M. Hayashida, Electrophoresis 22 (2001) 3785. [47] N. Ishizuka, H. Minakuchi, K. Nakanishi, N. Soga, H. Nagayama, Anal. Chem. 72 (2000) 1275. [48] Y. Shintani, X. Zhou, M. Furuno, H. Minakuchi, K. Nakanishi, J. Chromatogr. A 985 (2003) 351. [49] Y. Fan, Y.Q. Feng, S.L. Da, X.P. Gao, Analyst 129 (2004) 1065. [50] Y. Fan, Y.Q. Feng, S.L. Da, Z.G. Shi, Anal. Chim. Acta 523 (2004) 251. [51] Y. Fan, Y.Q. Feng, J.T. Zhang, S.L. Da, M. Zhang, J. Chromatogr. A 1074 (2005) 9. [52] J.C. Wu, J. Pawliszyn, Anal. Chem. 73 (2001) 55. [53] J. Wu, X. Yu, H. Lord, J. Pawliszyn, Analyst 125 (2000) 391. [54] J. Wu, J. Pawliszyn, J. Chromatogr. A 909 (2001) 37.

191

[55] H. Kataoka, Anal. Bioanal. Chem. 373 (2002) 31. [56] A. Mohammadi, Y. Yamini, N. Alizadeh, J. Chromatogr. A 1063 (2005) 1. [57] T.P. Gbatu, O. Ceylan, K.L. Sutton, J.F. Rubinson, A. Galal, A. Caruso, H.B. Mark, Anal. Commun. 36 (1999) 203. ¨ Ceylan, S. Oztemiz, ¨ [58] B.J. Yates, K.R. Temsamani, O. T.P. Gbatu, R.A. La Rue, U. Tamer, H.B. Mark Jr., Talanta 58 (2002) 739. [59] T. U˘gur, S. Y¨ucel, E. Nusret, U. Yasemin, P. Kadir, Y. Attila, J. Electroanal. Chem. 570 (2004) 6. [60] U. Tamer, N. Ertas¸, Y.A. Udum, Y. S¸ahin, K. Pekmez, A. Yıldız, Talanta 67 (2005) 245. [61] J. Wu, J. Pawliszyn, Anal. Chim. Acta 520 (2004) 257. [62] F. Guo, T. Gorecki, D. Irish, J. Pawliszyn, Anal. Commun. 33 (1996) 361. [63] H. Bagheri, A. Mir, E. Babanezhad, Anal. Chim. Acta 532 (2005) 89. [64] A. Ghassempour, N.M. Najafi, A. Mehdinia, S.S.H. Davarani, M. Fallahi, M. Nakhshab, J. Chromatogr. A 1078 (2005) 120. [65] H. Minjia, T. Chao, Z. Qunfang, J. Guibin, J. Chromatogr. A 1048 (2004) 257. [66] D. Djozan, S. Bahar, Chromatographia 59 (2004) 95. [67] M.A. Farajzadeh, N.A. Rahmani, Talanta 65 (2005) 700. [68] J.-F. Liu, N. Li, G.-B. Jiang, J.-M. Liu, J. J¨onsson, M.-J. Wen, J. Chromatogr. A 1066 (2005) 2. [69] T. Gorecki, P. Martos, J. Pawliszyn, Anal. Chem. 70 (1998) 19. [70] E.H.M. Koster, C. Crescenzi, W. Hoedt, K. Ensing, G.J. Jong, Anal. Chem. 73 (2001) 3140. [71] M. Farajzadeh, N.A. Rahmani, Anal. Sci. 20 (2004) 1359. [72] E.O. Otu, Pawliszyn, J. Mikrochim. Acta 112 (1993) 41. [73] J.-G. Hou, Q. Ma, X.-Z. Du, H.-L. Deng, J.-Z. Gao, Talanta 62 (2004) 241. [74] X.-Z. Du, Y.-R. Wang, X.-J. Tao, H.-L. Deng, Anal. Chim. Acta 543 (2005) 9. [75] Y. Liu, M.L. Lee, K.J. Hageman, Y. Yang, S.B. Hawthorne, Anal. Chem. 69 (1997) 5001. [76] D. Djozan, Y. Assadi, Chromatographia 45 (1997) 183. [77] F.M. Musteata, M. Walles, J. Pawliszyn, Anal. Chim. Acta 537 (2005) 231. [78] T.-H. Ding, H.-H. Lin, C.-W. Whang, J. Chromatogr. A 1062 (2005) 49. [79] C.L. Arthur, J. Pawliszyn, Anal. Chem. 62 (1990) 2145. [80] H.B. Wan, H. Chi, M.K. Wong, C.Y. Mok, Anal. Chim. Acta 298 (1994) 219. [81] C.P. Kuo, J. Shiea, Anal. Chem. 71 (1999) 4413. [82] D. Djozan, Y. Assadi, S.H. Haddadi, Anal. Chem. 73 (2001) 4054. [83] L. Pan, J. Pawliszyn, Anal. Chem. 69 (1997) 196. [84] C. Jurado, M.P. Gim´enez, T. Soriano, M. Men´endez, M. Repeto, J. Anal. Toxicol. 24 (2000) 11. [85] M.-K. Huang, C. Liu, S.-D. Huang, Analyst 127 (2002) 1203. [86] R. Herr´aez-Hern´andez, C. Ch´afer-Peric´as, P. Camp´ıns-Falc´o, Anal. Chim. Acta 513 (2004) 425. [87] M.-R. Lee, R.-J. Lee, Y.-W. Lin, C.-M. Chen, B.-H. Hwang, Anal. Chem. 70 (1998) 1963. [88] J. Carpinteiro, J.B. Quintana, I. Rodr´ıguez, A.M. Carro, R.A. Lorenzo, R. Cela, J. Chromatogr. A 1056 (2004) 179. [89] I. Rodr´ıguez, E. Rub´ı, R. Gonz´alez, J.B. Quintana, R. Cela, Anal. Chim. Acta 537 (2005) 259. [90] J.E. Hong, H. Pyo, S.-J. Park, W. Lee, Anal. Chim. Acta 539 (2005) 55. [91] J. Koziel, J. Noah, J. Pawliszyn, Environ. Sci. Technol. 35 (2001) 1481. [92] F. Reisen, S. Eschmann, R. Atkins, Environ. Sci. Technol. 37 (2003) 4664. [93] S.-W. Tsai, C.-M. Chang, J. Chromatogr. A 1015 (2003) 143. [94] P. Vesely, L. Lusk, G. Basarova, J. Seabrooks, D. Ryder, J. Agric. Food Chem. 51 (2003) 6941. [95] C. Deng, N. Li, X. Zhang, J. Chromatogr. B 813 (2004) 47. [96] C. Deng, J. Zhang, X. Yu, W. Zhang, X. Zhang, J. Chromatogr. B 810 (2004) 269. [97] Q. Wang, J. O’Reilly, J. Pawliszyn, J. Chromatogr. A 1071 (2005) 147.

192

C. Dietz et al. / J. Chromatogr. A 1103 (2006) 183–192

[98] E.E. Stashenko, M.A. Puertas, W. Salgar, W. Delgado, J.R. Martınez, J. Chromatogr. A 886 (2000) 175. [99] E.E. Stashenko, J.R. Martinez, Trends Anal. Chem. 23 (2004) 553. [100] H. Liu, P.K. Dasgupta, Anal. Chem. 68 (1996) 1817. [101] M.A. Jeannot, F.F. Cantwell, Anal. Chem. 68 (1996) 2236. [102] E. Psillakis, N. Kalogerakis, Trends Anal. Chem. 21 (2002) 53. [103] D.A. Lambropoulou, E. Psillakis, T.A. Albanis, N. Kalogerakis, Anal. Chim. Acta 516 (2004) 205. [104] E. Psillakis, N. Kalogerakis, J. Chromatogr. A 938 (2001) 113. [105] L. Vidal, A. Canals, N. Kalogerakis, E. Psillakis, J. Chromatogr. A 1098 (2005) 25. [106] Y.C. Fiamegos, C.G. Nanos, C.D. Stalikas, J. Chromatogr. B 813 (2004) 89. [107] M.C. Lopez-Blanco, S. Blanco-Cid, B. Cancho-Grande, J. SimalGandara, J. Chromatogr. A 984 (2003) 245. [108] P. Das, M. Gupta, A. Jain, K.K. Verma, J. Chromatogr. A 1023 (2004) 33. [109] D.A. Lambropoulou, T.A. Albanis, J. Chromatogr. A 1049 (2004) 17. [110] P. Liang, L. Guo, Y. Liu, S. Liu, T.-Z. Zhang, Microchim. J. 80 (2005) 19. [111] R. Batlle, C. Ner´ın, J. Chromatogr. A 1045 (2004) 29. [112] C. Deng, N. Yao, A. Wang, X. Zhang, Anal. Chim. Acta 536 (2005) 237. [113] C. Deng, X. Yang, X. Zhang, Talanta 68 (2005) 6. [114] Y. Yamini, M. Hojjati, M. Haji-Hosseini, M. Shamsipur, Talanta 62 (2004) 265. [115] M. Saraji, J. Chromatogr. A 1062 (2005) 15. [116] V. Colombini, C. Bancon-Montigny, L. Yang, P. Maxwell, R.E. Sturgeon, Z. Mester, Talanta 63 (2004) 555. [117] S. Gil, S. Fragueiro, I. Lavilla, C. Bendicho, Spectrochim. Acta Part B 60 (2005) 145. [118] S. Fragueiro, I. Lavilla, C. Bendicho, Spectrochim. Acta Part B 59 (2004) 851. [119] M.H. Ma, F.F. Cantwell, Anal. Chem. 71 (1999) 388. [120] Y. Li, T. Zhang, P. Liang, Anal. Chim. Acta 536 (2005) 245. [121] J.-F. Liu, Y.-G. Chi, G.-B. Jiang, C. Tai, J.-F. Peng, J.-T. Hu, J. Chromatogr. A 1026 (2004) 143. [122] J.-F. Liu, J.-F. Peng, Y.-G. Chi, G.-B. Jiang, Talanta 65 (2005) 705. [123] E. Psillakis, N. Kalogerakis, Trends Anal. Chem. 22 (2003) 565. [124] C. Basheer, R. Balasubramanian, H.K. Lee, J. Chromatogr. A 1016 (2004) 11. [125] M. Charalabaki, E. Psillakis, D. Mantzavinos, N. Kalogerakis, Chemosphere 60 (2005) 690. [126] L. Hou, H.K. Lee, J. Chromatogr. A 1038 (2004) 37. [127] F.J.M. de Santana, A.R. Moraes de Oliveira, P.S. Bonato, Anal. Chim. Acta 549 (2005) 96.

[128] [129] [130] [131] [132] [133] [134] [135] [136] [137] [138] [139] [140] [141] [142] [143] [144] [145] [146] [147] [148] [149] [150] [151] [152] [153] [154] [155] [156]

C. Basheer, J.P. Obbard, H.K. Lee, J. Chromatogr. A 1068 (2005) 221. D.A. Lambropoulou, T.A. Albanis, J. Chromatogr. A 1061 (2004) 11. H.-J. Pan, W.-H. Ho, Anal. Chim. Acta 527 (2004) 61. C. Basheer, H.K. Lee, J. Chromatogr. A 1057 (2004) 163. E. Psillakis, D. Mantzavinos, N. Kalogerakis, Anal. Chim. Acta 501 (2004) 3. C. Basheer, H.K. Lee, J.P. Obbard, J. Chromatogr. A 1022 (2004) 161. J.B. Quintana, R. Rodil, T. Reemtsma, J. Chromatogr. A 1061 (2004) 19. S. Pedersen-Bjergaard, K.E. Rasmussen, Anal. Chem. 71 (1999) 2650. M. Marlow, R.J. Hurtubise, Anal. Chim. Acta 526 (2004) 41. E. Gonz´alez-Pe˜nas, C. Leache, M. Viscarret, A. P´erez de Obanos, C. Aragu´as, A. L´opez de Cerain, J. Chromatogr. A 1025 (2004) 163. H.G. Ugland, M. Krogh, L. Reubsaet, J. Chromatogr. B 798 (2003) 127. A.S. Yazdi, Z. Es’haghi, Talanta 66 (2005) 664. T.S. Ho, J.L.E. Reubsaet, H.S. Anthonsen, S. Pedersen-Bjergaard, K.E. Rasmussen, J. Chromatogr. A 1072 (2005) 29. A.S. Yazdi, Z. Es’haghi, J. Chromatogr. A 1082 (2005) 136. J.-X. Wang, D.-Q. Jiang, X.-P. Yan, Talanta (2005) in press (Available online 21 July 2005). C. Basheer, V. Suresh, R. Renu, H.K. Lee, J. Chromatogr. A 1033 (2004) 213. E. Psillakis, N. Kalogerakis, Trends Anal. Chem. 22 (2003) 565. S. Pedersen-Bjergaard, K.E. Rasmussen, J. Chromatogr. B 817 (2005) 3. K.E. Rasmussen, S. Pedersen-Bjergaard, Trends Anal. Chem. 23 (2004) 1. C. Blasco, M. Fern´andez, Y. Pic´o, G. Font, J. Chromatogr. A 1030 (2004) 77. H.A. Soini1, K.E. Bruce1, D. Wiesler, F. David, P. Sandra, M.V. Novotny, J. Chem. Ecol. 31 (2005) 1573. W. Liu, H. Wang, Y. Guan, J. Chromatogr. A 1045 (2004) 15. J. Pettersson, A. Kloskowski, C. Zaniol, J. Roeraade, J. Chromatogr. A 1033 (2004) 339. L. Wang, A. Hosaka, C. Watanabe, H. Ohtani, S. Tsuge, J. Chromatogr. A 1035 (2004) 277. D. Sulistyorini, M.D. Burford, D.J. Weston, D.W.M. Arrigan, Anal. Lett. 35 (2002) 1429. J.-P. Lambert, W.M. Mullett, E. Kwong, D. Lubda, J. Chromatogr. A 1075 (2005) 43. I. Bruheim, X.C. Liu, J. Pawliszyn, Anal. Chem. 75 (2003) 1002. Y. Hu, Y. Yang, J. Huang, G. Li, Anal. Chim. Acta 543 (2005) 17. C. Basheer, A. Parthiban, A. Jayaraman, H.K. Lee, S. Valiyaveettil, J. Chromatogr. A 1087 (2005) 274.