Anal Bioanal Chem (2006) 384: 1447–1461DOI 10.1007/s00216-005-0242-z

REVIEW

José Benito Quintana . Isaac Rodríguez

Strategies for the microextraction of polar organiccontaminants in water samples

Received: 4 October 2005 / Revised: 14 November 2005 / Accepted: 18 November 2005 / Published online: 22 February 2006# Springer-Verlag 2006

Abstract In this paper the most recent developments in themicroextraction of polar analytes from aqueous environ-mental samples are critically reviewed. The particularitiesof different microextraction approaches, mainly solid-phase microextraction (SPME), stir-bar-sorptive extraction(SBSE), and liquid-phase microextraction (LPME), andtheir suitability for use in combination with chromato-graphic or electrically driven separation techniques fordetermination of polar species are discussed. The compat-ibility of microextraction techniques, especially SPME,with different derivatisation strategies enabling GC deter-mination of polar analytes and improving their extractabil-ity is revised. In addition to the use of derivatisationreactions, the possibility of enhancing the yield of solid-phase microextraction methods for polar analytes by usingnew coatings and/or larger amounts of sorbent is alsoconsidered. Finally, attention is also focussed on describingthe versatility of LPME in its different possible formats andits ability to improve selectivity in the extraction of polaranalytes with acid-base properties by using separationmembranes and buffer solutions, instead of organicsolvents, as the acceptor solution.

Keywords Solid-phase microextraction . Liquid-phasemicroextraction . Single-drop microextraction . Membraneextraction . Stir-bar-sorptive extraction . Polar organiccontaminants . Water analysis

Introduction

Until the mid-90s, organic trace analysis of water wasmainly focussed on persistent organic pollutants (POP)such as PCB, PAH, organochlorine pesticides, etc. Some ofthese are highly hydrophobic compounds, easily bioaccu-mulated and biomagnified through the trophic chain in theaquatic system. Fortunately, nowadays, most of these havebeen banned and their environmental concentrations arestrictly controlled. There is now, in contrast, increasinginterest on the fate and role of polar organic contaminantsin the aqueous environment. Many of these compounds areemployed as household chemicals. Several pharmaceuticaldrugs, disinfection agents, pesticides, and different person-al care products can be included in this group.

Although, in general, polar species are not bio-accumulative, some are resistant to degradation duringconventional wastewater treatment. This, and their ex-cellent water solubility and continuous discharge into theenvironment, facilitates the ubiquitous distribution of somepolar species in aquatic media. As an example, EDTA hasbeen found to be the most abundant anthropogenic organiccompound in German surface waters [1]. Surface water isused very often as a source of drinking water. As aconsequence, concentrations of EDTA between 1 and 5 μgL−1 have been reported for tap water in Germany [1].

In parallel with these findings analytical chemistry alsois evolving toward more environmentally responsiblepolicies, searching for sample-preparation approacheswith lower consumption of toxic organic solvents tominimise the generation of hazardous residues and healthrisks for operators. This was partially achieved in the firstinstance by the commercialisation of SPE, which replacedLLE, and was further improved by the popularisation ofSPME [2, 3], a completely solvent free technique.

The development of SPME has been followed by othermicroextraction techniques, for example stir-bar-sorptiveextraction (SBSE) [4], liquid-phase microextraction(LPME) [5−7], and membrane extraction techniques [8,9]. All share a common idea—analytes are extracted intoan acceptor medium (sorbent or solution) by equilibrium

J. B. Quintana (*)Department of Water Quality Control,Technical University of Berlin,Sekr KF 4, Strasse des 17 Juni 135,10623 Berlin, Germanye-mail: sttito@usc.esTel.: +49-30-31424281Fax: +49-30-31423850

J. B. Quintana . I. RodríguezDepartamento de Química Analítica, Nutrición e Bromatoloxía,Facultade de Química, Instituto de Investigación e AnáliseAlimentario, Universidade de Santiago de Compostela,15782 Santiago de Compostela, Spain

processes. Thus, in microextraction techniques only afraction of the analytes contained in the sample isrecovered, in contrast with SPE and LLE which areexhaustive processes. In summary, solvent-free extractiontechniques are more sustainable and easily implemented,and also reduce exposure of the analyst to solvents. Apartfrom these advantages they enable more selectivity insample preparation than exhaustive extraction approachessuch as SPE or LLE.

The objective of this paper is to critically review somekey features of microextraction strategies suitable foranalysis of polar organic compounds in water, includingthose aspects that require further investigation and/or maybe subject to important development in the future. Thetarget analytes considered are medium to high polarityspecies—pharmaceuticals, personal care products, pesti-cides, disinfection by-products, amines, and phenoliccompounds, etc.

Solid-phase microextraction (SPME)

Although SPME, especially in combination with GC, is awell established sample-preparation technique, its applica-tion to the determination of polar analytes in water samplesis still an emerging field. Although PA and CW-DVBcoatings are commercially available, extraction efficiencyfor highly polar analytes is still limited and the develop-ment of more polar coatings is of interest. Some otherdifficulties of this type of analysis are the need forderivatisation before GC determination and problemsassociated with LC determination, for example the stabilityof some coatings when exposed to organic solvents andpeak broadening during on-line desorption of the analytesin the SPME-LC desorption interface.

These three aspects of SPME are considered separatelyin the following paragraphs. Basic principles of thetechnique, and general guidelines, are not discussedbecause they have been discussed in detail in severalbooks and reviews (e.g. [2, 3, 10]).

Development of new coatings

In recent years growing interest in the determination ofpolar compounds has fostered the development of newsorbents for solid-phase extraction (SPE), as recentlyreviewed [11]. This has been achieved by different researchgroups and also by several private companies, leading tomuch competence. For SPME, however, the situation isquite different—there is no such competence commerciallyand, thus, only research groups are currently developingdifferent sorbents for incorporation into microextractionfibres. Some of these are specifically intended for thedetermination of polar analytes in water samples. The mostrecent research on different sorbents is considered here,with emphasis on possible future developments towardanalysis of polar compounds.

The first and probably most important development hasbeen the use of sol-gel technology for coating the silicafibres. The pioneering work was performed by Chong et al.[12], who used this method to prepare sol-gel polydi-methylsiloxane (PDMS) fibres. Unlike commercial PDMSfibres the sol-gel variety have the PDMS chemicallybonded to the silica core, which increases their thermalstability and surface area by creation of a highly cross-linked network. Titanium and zirconium-based materialshave also been prepared recently; these have increased thepH and mechanical stability of SPME fibres [13, 14].Similarly, a new generation of PDMS fibres based on a Zralloy core will soon be available commercially fromSupelco; these are expected to enhance both extractionefficiency for very acidic and basic compounds, byenabling work with samples adjusted to extremes of pH,and the mechanical stability of the fibre itself, which isespecially important when the SPME holder is incorpo-rated in an autosampler device for GC.

This sol-gel technology has been further used to preparepoly(ethylene glycol) (PEG) [15], PDMS/DVB [16],PDMS/poly(vinyl alcohol) (PDMS/PVA) [17], and poly-tetrahydrofuran (PTHF) [18] coated fibres. All of these,with the exception of PEG, are relatively apolar polymers.

More polar sol-gel bonded phases have been preparedfrom crown ethers [19−22] and calyx[4]arenes [23, 24].Use of aromatic crown ethers in the sol-gel network hasenabled more efficient HS-SPME of aromatic amines(compared with use of commercial PA fibres), due to π-π,dipole, and H-bonding interactions [22]. Similar resultswere also obtained in the concentration of amines by use ofan amide-modified calyx[4]arene sorbent [24]. The polar-ity of the generated coating can be tuned by selectingappropriate monomers for polymerisation. An example ofthis is the synthesis of a polyacrylate fibre for extraction of2-chloroethyl ethyl sulfide from soil [25]. In that work,three different monomers were considered: methyl acry-late, methyl methacrylate and butyl methacrylate. The bestresults were obtained by use of butyl methacrylate (BMA),because both analyte and extracting sorbent are of mediumpolarity and similar log Kow values. The same monomer(BMA) can be sol-gel copolymerised with DVB, resultingin a polar coating with enhanced surface area and highextraction efficiency, compared with commercial PA andPDMS-DVB fibres, in the determination of polar alcoholsand acids in wine samples [26]. By a method similar to thatused for BMA-based coatings, Basheer et al. [27] preparedSPME sol-gel fibres coated with oligomeric bisphenolicgroups, the hydrophobicity of which is determined by theratio of ether to free phenolic groups.

Inorganic materials, for example anodised metal wires(e.g. CuCl), which can participate in ionic interactions,have also been tested as SPME fibres for concentration ofamines [28, 29]. Thermal stability, ease preparation, andrelatively low cost are their main advantages. Thesesorbents are deactivated by water, however, so can beused only with gaseous samples or in headspace (HS)mode, at room temperature, for water samples. These are

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rather unfavourable conditions for extraction of polaranalytes with high water solubility and low volatility.

The third group of materials prepared for SPME of polarcompounds are electropolymerised polypyrrole (PPy),polyphenylpyrrole (PPPy) [30−32], and polyaniline [33].

The last of these has been used for SPME of phenols fromwater samples and PPy and PPPy have been used for in-tube SPME of basic compounds from water and biologicalsamples, for which they performed better than commer-cially available polar PEG capillaries [30, 31]. PPy and

Table 1 Overview of different approaches for combining SPME and derivatisation for analysis of polar organic contaminants in water

Analytes Sampling modea Type of derivatisation Fibre type Derivatisation reagentb Refs.

Aldehydes and ketones DS In-sample PDMS PFBHA [43]HS In-sample PDMS and

PDMS-DVBPFBHA [43, 44]

HS On-fibre duringextraction

PDMS-DVB PFBHA [45]

Alkylthiols DS In-sample PDMS-DVB DNFB [46]DS In-sample PDMS-DVB NEM [47]

Short-chain fatty acids HS In-sample PA PFBBr/PFPDE [49]HS On-fibre during

extractionPA PDAM [49, 124]

Acetic and haloacetic acids HS In-sample PA Benzyl bromide [48]HS In-sample PDMS H2SO4/EtOH [51]HS In-sample CAR-PDMS DMS [52]HS In-sample PDMS HCl/MeOH [50]

Nonylphenol ethoxylatesand carboxylates

HS In-sample CW-DVB DMS [125]

Long-chain fatty acids DS On-fibre after extraction PDMS Diazomethane [49]DS In-port PA TMAOH/TMAHS [49]

Acidic herbicides DS In-sample PDMS Butyl chloroformate [54]DS In-sample PDMS-DVB Benzyl bromide [53]HS On-fibre during

extractionCW-DVB PFBBr [53]

DS On-fibre after extraction PA Diazomethane [55]DS On-fibre after extraction PA MTBSTFA [58]

Phthalic acid monoesters DS On-fibre after extraction PDMS-DVB Diazomethane [56]Acidic drugs DS On-fibre after extraction PA MTBSTFA [40]Degradation productsof warfare agents

DS On-fibre after extraction CAR-PDMS MTBSTFA [57]

Oestrogens and anabolic steroids DS On-fibre after extraction PA and CW-DVB BSTFA [64]DS On-fibre after extraction PA MSTFA [39]

Phenolic compounds DS On-fibre after extraction PA and PDMS-DVB MTBSTFA [61]DS In-sample PA Acetic anhydride [62]HS In-sample CAR-PDMS and PDMS Acetic anhydride [63]

Linear alkylbenzensulfonates DS In-port PDMS TMAHS [59]Perfluorocarboxylic acids DS In-port PDMS TMAHS [60]Basic drugs DS In-sample PDMS-DVB Acetic anhydride [65]Aromatic amines DS In-sample PDMS-DVB NaNO2/hydroiodic acid [68]Aliphatic amines HS In-sample Crown ether sol-gel fibre TFBza-suc [22]

HS In-sample PDMS SIBA [67]HS In-sample PA PFBAY [66]

aHS: headspace sampling; DS: direct samplingbPFBHA: pentafluorobenzylhydroxylamine; DNFB: 2,4-dinitrofluorobenzene; NEM: N-ethylmaleimide; PFBBr: pentafluorobenzylbromide; PFPDE: pentafluorophenyldiazoethane; PDAM: pyrenyldiazomethane; DMS: dimethylsulfate; TMAOH: tetramethylammoniumhydroxide ; TMAHS: tetramethylamonium hydrogen sulfate; MTBSTFA: N-methyl-N-(tert-butyldimethylsilyl)trifluoroacetamide; BSTFA:bis-(trimethylsilyl)trifluoroacetamide; MSTFA: N-methyl-N-(trimethylsilyl)trifluoroacetamide; TFBza-suc: tetrafluorobenzoic acidN-hydroxysuccinimide ester; SIBA: N-succinimidyl benzoate; PFBAY: pentafluorobenzyl aldehyde

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PPPy are cationic sorbents, but their selectivity can betuned by choosing an appropriate counterion. If this is, forexample,ClO�

4 ; they have some anion-exchange character,whereas this behaviour is lost when bigger anions (e.g.dodecyl sulfate) are used, resulting in more hydrophobicsorbents [32]. Similarly, the selectivity of aniline polymersis determined by their redox state. The less oxidisedpolymers produce the best results for polar analytes, byenabling hydrogen-bonding interactions through –NHgroups [34]. A drawback of these electropolymerisedmaterials is, unfortunately, their thermal instability above200°C; they are, therefore, best suited to in-tube SPME andsolvent desorption, rather than thermal, for example GC,applications. This limitation might be overcome by use ofroom-temperature-ionic liquids as SPME coatings. Thesesolvents are virtually non-volatile and their selectivitytoward polar analytes can be modified by choosing theappropriate anion and cation components. Preliminaryresearch on SPME with ionic liquids has been conductedby Liu et al. [35]. In their work 1-octyl-3-methylimidazol-ium hexafluorophosphate ([C8MIM][PF6]) was physicallycoated on a silica fibre by dipping it in this solvent. Its highviscosity and thermal stability enabled HS-SPME of BTEXfrom paints and their determination by GC. It is expectedthat further applications can be extended to polar analytesby the appropriate selection of both the cation and anion, ina way similar to that used with PPy polymers. Ionic liquidshave already been evaluated as stationary phases in GCcolumns and shown to have excellent thermal stability andthe ability to separate polar and non-polar compounds [36].

Derivatisation

Despite the advances in the development of new SPMEsorbents described above, derivatisation of polar analytes isstill important, because it enables their hydrophobicity tobe increased, which normally leads to higher extractionyields when using commercially available SPME fibres,and improvement of performance when SPME is combinedwith GC analysis. These benefits are clear when detectionlimits of applications using SPME and GC for determina-tion of polar compounds [37, 38] are compared with valuesachieved after incorporation of an additional derivatisationstep in the analytical procedure [39, 40]. This aspect has,however, received little attention and, for example, in areview from 2003 on SPME of herbicides in environmen-tal samples [41], it was not even considered, even thoughsome of the target species were difficult to determine byGC without derivatisation. Only recently the combinationof SPME and derivatisation has been reviewed in a moregeneral context [42]. Our review focuses on water analy-sis and considers the most recent applications andachievements.

Table 1 summarizes different applications of SPME-derivatisation in gas chromatographic analysis of polarorganic compounds in water. Different strategies can beused for sample, on-fibre, and in-port derivatisation.

The first is also the simplest. Derivatisation occurs in theaqueous sample before, or simultaneously with, theextraction step. It improves both the affinity of the parentanalytes for the fibre and the efficiency of subsequent GCseparation. For obvious reasons this strategy is not suitablefor moisture-sensitive reagents; these are, however, com-patible with on-fibre derivatisation reactions. On-fibrederivatisation can be performed either by preloading thefibre with the derivatisation agent, so the reaction occurs assoon as analytes are incorporated in the sorbent material(for water-sensitive reagents only the HS mode can beemployed) or, alternatively, by first concentrating theanalytes in the fibre and then exposing the fibre to thevapour of the derivatisation reagent. In the last strategy (in-port derivatisation) polar analytes, with acid-base proper-ties, are extracted in the SPME fibre as ion pairs which arefurther decomposed, at the high temperatures of the GCinjection port, to produce volatile by-products and the alkylderivatives of the target compounds. The most suitableapproach depends on the properties of the analytes and thederivatisation reaction to be performed.

The first example considered (Table 1) is the determi-nation of volatile compounds such as aldehydes, ketones,and thiols. Although some of these are neutral and volatilespecies, because of their reactivity and polarity they mustbe derivatised to increase their affinity for the SPME fibreand their thermal stability for GC analysis. Aldehydes andketones can be converted into oximes by reaction withpentafluorobenzylhydroxylamine (PFBHA), either directlyin the sample [43, 44] or in a SPME fibre previously loadedwith PFBHA, using the HS mode [45]. Experimentalresults showed that the second option was faster than thefirst [45]. For determination of thiols, however, in-samplederivatisation is preferred and two derivatisation agentshave been explored—2,4-dinitrofluorobenzene (DNFB)[46], which introduces appropriate groups for ECD andNPD determination, and N-ethylmaleimide (NEM), whichis regarded as being specific for thiols [47]. Whereasserious problems related to matrix effects have beenencountered when using DNFB, it seems that reactionwith NEM proceeds faster and without such difficulties.

For species containing carboxyl groups different deriv-atisation approaches have been considered, depending onmolecular weight, pKa values and polarity. The use of alkylhalides [48, 49] or strongly acidic conditions [50−52]enables esterification of carboxyl compounds in watersamples. Further SPME must then be performed in HSmode to avoid damage of the coated phase. This strategy isvalid for low-molecular-weight analytes but leads to highdetection limits for heavier compounds with low vapourpressures, even after derivatisation [53, 54]. This problemcan be overcome by first extracting the compounds bydirect exposure of the SPME fibre to the sample and thenperforming on-fibre derivatisation; this enables the use ofmoisture-sensitive reagents such as diazomethane [55, 56]and silylation agents [40, 57, 58]. In this option sample pHmust be adjusted to acidic values to obtain the compoundsin their neutral forms. Under these conditions the highestextraction yields are usually obtained with the PA fibre.

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Problems arise for non-volatile species with pKa valuesbelow 1 unit. For such species, addition of an ion-pairingagent to the water sample adjusted to a pH above the pKa ofthe analytes, then direct SPME, is the most convenientsolution. The ion-pairing agent also acts as an alkylatingagent in the injection port of the chromatograph [59, 60].

For less acidic analytes, e.g. phenols, both on-fibre afterextraction [61] and in-sample [62, 63] derivatisationprocedures can be employed. The acylation reaction usedfor derivatisation of these compounds in aqueous media,normally using acetic anhydride, occurs under milderconditions (compared with esterification of carboxylicacids) and, therefore, both headspace and direct SPMEapproaches are possible. The derivatives obtained are,moreover, more hydrophobic and volatile, which alsoimproves their extraction yield in headspace sampling.Acylation of aliphatic alcohol groups is not possible,however, and for these analytes on-fibre silylation ispreferred [39, 64]. So, for example, the on-fibre derivatisa-tion of oestrogens with N-methyl-N-(trimethylsilyl)trifluo-roacetamide (MSTFA) enables their determination at lowng L−1 levels in wastewater samples (Fig. 1) [39].

The last group of compounds presented in Table 1 arebasic analytes. In-sample acylation [65] or derivatisationwith pentafluorobenzyl aldehyde (PFBAY) [66] are alsopossible for some amines, although some authorsdecided to investigate specifically designed reagents[22, 67, 68]. Another possibility, which has been used forthe determination of amphetamine drugs, entails on-fibrederivatisation of the analytes while they are beingheadspace-extracted, with the particularity that an insertis used to support the derivatising agent (HFBCl-HFBA)in the HS over the sample (Fig. 2) [69, 70]. This insertcontains several holes enabling the analytes to enter it.The SPME fibre is therefore exposed simultaneously totarget species and vapour from the derivatisation reagent,but not to the water sample.

In-tube SPME

In contrast with the successful combination of SPME andGC, tandem SPME-HPLC has been less extensively used.This is mainly attributed to two shortcomings—the lack ofa wide range of polar coatings (now partially overcome)and problems related to the commercial desorption cham-ber [71, 72], which often led to peak broadening. Althoughthe design of the desorption chamber has been improved byreducing its internal volume [71], some authors concludedthat the best solution is simply to perform an off-linedesorption of the SPME fibre with a smaller volume ofsolvent, injecting only part of the extract into the HPLCsystem [73]. Obviously, as a result, sensitivity is reducedand automation capability is lost.

Another drawback of SPME which has encouraged thedevelopment of in-tube SPME was the limited availabilityof LC solvent-compatible coatings. Even the robust PApolymer, employed in SPME for concentration of polarspecies, deteriorates when exposed to organic solvents,compromising the extraction of polar compounds. In-tubeSPME overcomes this limitation by using commerciallyavailable GC capillary columns which are (thermally andchemically) resistant and available in a wide range ofpolarities [74].

When combined with HPLC, in-tube SPME alsofacilitates automation of the sample-preparation step [75].Most forms of on-line coupling of in-tube SPME and LCare based on simple modification of the HP-1100 seriesautosampler (Agilent Technologies) by installing a GCcapillary between the injection loop and the needle (Fig. 3).The original HPChem software can be used to operate thewhole system automatically. Diffusion of the analytes fromthe sample to the sorbent phase is speeded up by aspiratingand ejecting it several times through the capillary. Inpractice, however, equilibrium is almost never achieved ina reasonable time, then non-equilibrium conditions areemployed. As an example, Fig. 4a shows the extractionprofile obtained from five oestrogenic compounds ex-

Fig. 1 GC-MS-MS chromato-gram obtained from oestrone(8 ng L−1) and oestradiol (5 ngL−1) in an (unspiked) waste-water effluent after SPME andon-fibre derivatisation withMSTFA. Reproduced from Ref.[39], copyright (2004), withpermission from Elsevier

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tracted from water; even after 20 draw/eject cyclesequilibrium is not reached. The efficiency of the processcannot, furthermore, be improved by increasing the draw/eject flow rate, because above 100 μL min−1 there is areduction on the extraction yield (Fig. 4b) because of theformation of bubbles in the capillary [76]. Finally, afterextraction, the compounds are desorbed by the mobilephase, or by drawing a small amount of a solvent from adifferent vial [75], and analysed by LC.

Of commercially available GC coated capillaries, poly(ethylene glycol) coatings (Omegawax, DB-WAX, etc.)have resulted in the best extraction efficiency in theanalysis of polar contaminants in aqueous samples(Table 2). Use of the adsorptive coated capillary Supel-Q-PLOT (DVB polymeric material) has recently beenproved to be more efficient for analysis of oestrogens,because of its large surface area, which improves mass-

transfer kinetics [76], and because oestrogens are ofintermediate polarity.

Despite the wide availability of coatings and thepossibility of automation of this technique, the number ofapplications to analysis of polar compounds in water by in-tube SPME-LC is rather small (Table 2). This is probablybecause, normally, the sample volume considered is limitedto 1 mL and the extraction efficiency to a maximum of ca.30%. As a result, the concentration factors achieved can beregarded as appropriate for biological samples but may notbe sufficient for environmental analysis. Obviously, anadvantage of in-tube SPME that can be exploited is theselectivity, if performed under appropriate conditions. Thisis still a major problem for analysis of complex matricessuch as wastewater. Even when LC-MS-MS is used as thedetermination technique, accuracy and detection limits ofthe method are seriously impaired, because of the presence

Fig. 2 Schematic diagram ofsimultaneous SPME and on-fibre derivatisation of amphet-amine compounds. Reproducedfrom Ref. [70], copyright(2005), with permissionfrom Elsevier

Fig. 3 Schematic diagramof an in-tube SPME-LC-MSsystem [75]

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of large amounts of co-extracted species which disturb theefficiency of the electrospray ionisation step [77]. Thisaspect has not yet been considered in published articles,which focus mostly on maximising the extractionefficiency.

This objective can be achieved either by increasing theamount of sample or by increasing the coating/sampleratio. The first option is not normally adopted, because ofincompatibilities with some autosampler configurationsand the consequent increase in equilibration time. Thesecond can be accomplished in several ways. The simplestis to increase the coating thickness and/or the length of theGC capillary; this, however, has the disadvantage of slowerkinetics during both sorption and desorption [75]. Anotherpossibility is the so called wire-in-tube SPME [71]. Thisentails introducing a narrow wire into the coated capillaryso that the sample coating/sample ratio is doubled; a furtherdevelopment of this is to pack the capillary with filamentsof a polymeric material ( fibre-in-tube) [71] or with amonolithic sorbent [78]. The last option is the mostpromising. It operates as a chromatographic enrichmentprocess and not as an equilibrium technique and can beregarded, therefore, as miniaturised SPE rather than SPME.Most of the sorbents employed are quite hydrophobic andhave not been tested for extraction of polar analytes.

GC capillaries can also be used to perform miniaturisedSPE. The analytes can then be desorbed by use of anappropriate solvent [79], or thermally [80], makingcoupling to GC easier. Although described as in-tubeSPME [79, 80] or capillary microextraction [15, 18] by

most authors, it must not be forgotten that this isminiaturised SPE and, therefore, maximum enrichment isdetermined by breakthrough of the analytes and not solelyby kinetic or equilibrium processes [15, 18].

Stir-bar sorptive extraction (SBSE)

SBSE was developed at the “Research Institute of Chro-matography” (Kortrijk, Belgium) [4] and commercialisedby Gerstel (Mülheim, Germany) in 1999 under the name“Twister”. Analytes are concentrated using a magnetic stir-bar coated with PDMS which is placed in the liquidsample. This format results in a significant increase in thevolume of the extraction phase—from approximately0.5 μL for an SPME fibre (100 μm PDMS) to ca 126 μLfor SBSE, although the stir-bars most commonly used(10 mm length, 0.5 mm coating thickness) have a PDMSvolume of ca 24 μL, which is still a 50-fold increase. As aconsequence, the yield of the extraction process is muchgreater when using a stir-bar rather than an SPME fibre,both coated with PDMS. This effect is even more importantfor species of medium polarity than for very hydrophobicand very hydrophilic species [4]. The greater coating areaof magnetic stir bars does, on the other hand, alsosomewhat limit the applicability of the technique—theextraction kinetics are slower than for SPME fibres, athermal desorption unit is necessary for transfer of theanalytes from the Twister bars to the head of the GCcolumn, and very few publications describe its coupling to

Fig. 4 Effects of (a) draw/ejectcycle and (b) flow rate onthe in-tube SPME of oestrogens.Reprinted from Ref. [76],copyright (2005), withpermission from Elsevier

Table 2 Applications of in-tube SPME combined with HPLC for determination of polar analytes in water samples

Analyte Capillary coating Detection Refs.

Phenylurea herbicides Omegawax, SBP-5, PPya, PMPya LC-UV, LC-MS [126, 127]Phenoxy acid herbicides DB-WAX LC-MS [128]Carbamate pesticides Omegawax 250 LC-UV, μLC-UV [74, 129, 130]Phenols, β-blockers, aromatic compounds PPya LC-UV [31, 131]Oestrogens Supel-Q-PLOT LC-MS-MS [76]Organoarsenic species PPya LC-MS [132]aNon-commercial coatings: PPy: polypyrrole; PMPy: polymethylpyrrole

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LC, because it requires relatively large solvent volumes fordesorption and, obviously, on-line connection of bothtechniques is not possible.

Because of the apolar character of PDMS, SBSE hasmainly been successfully applied to the determination ofmedium to non-polar pesticides [81, 82]; it fails in theextraction of more polar compounds [82, 83] unless theyhave previously been derivatised. As in SPME, both in-sample acylation (normally acetylation) and in-port silyla-tion strategies can be considered. In-sample acetylation canbe used for extraction of analytes containing phenolicmoieties: e.g. hydroxy-PAH [84], chlorophenols andalkylphenols [85−89], and oestrogens [90]. In-port silyla-tion has been used for determination of alkylphenols [91]by incorporating a small glass capillary tube containing thederivatising agent (0.5 μL BSTFA) with the PDMS-coatedstir bar in the desorption chamber. The advantage of thisreaction over acylation is that in-port silylation can be usedto derivatise a wider range of functional groups and notonly phenol moieties; acylation, on the other hand, enablesthe Kow of the analyte to be increased, improving theextraction yield. Both approaches can be combined toimprove sensitivity in the determination of compoundscontaining aromatic hydroxyl and other polar groups whichreact with silylation reagents. This has been applied tothe determination of oestradiol [92]—after acetylation ofthe phenolic moiety the affinity of the hormone for thePDMS polymer increases; silylation of the aliphatichydroxyl during desorption of the stir bar further improvesthe peak shape and thus the minimum amount detectable byGC-MS.

In-sample derivatisation of polar analytes to producemore hydrophobic species is not always possible, however,and for the most polar compounds extraction is stilldifficult with PDMS-coated stir bars [82, 83]. To overcomethis limitation a new dual-phase stir bar has been producedand commercialized by Gerstel. It consists of an emptyPDMS tube (1 cm long, 0.5-1 mm thick) which can befilled with the desired sorbent (Fig. 5). Only carbonaceousmaterials have yet been tested as filling sorbents. Theresults obtained revealed important enhancement of theextraction yield for polar compounds (flavour compoundsand pesticides) compared with those achieved using theempty PDMS tube [93]. Although additional studies arestill necessary to assess the durability of this device, cost ofextraction, and potential carry-over problems for lessvolatile species, because of the high retention capacity ofcarbonaceous sorbents, it is expected this dual-phase

configuration will help to increase the range of applicationsof the SBSE technique.

An interesting alternative to commercial stir bars (i.e.Twister), based on the use of disposable PDMS-coatedrods, has been proposed by Montero, Popp and coworkers[94, 95]. These rods have the advantage that they can be cutto the desired length in the laboratory. These rods areavailable in diameters from 1 to 5 mm. This determines theratio sorbent volume/sorbent amount and thus the kineticsof the extraction process. Because they are very inexpen-sive, they can be discarded after each use, reducing thepossibility of cross-contamination between samples andthe need for cleaning. Although such rods have been usedfor determination of hydrophobic analytes only (PCB,chlorobenzenes, and PAH), it seems reasonable that asimilar strategy could be employed for more polarcompounds by selection of more polar materials.

Liquid-phase and membrane microextraction

The last technique considered in this review is liquid-phasemicroextraction (LPME), which is the result of applyingthe principles of SPME, i.e. reduced organic solventconsumption and equilibrium extraction, to LLE. Thereare, in practice, several means of achieving theseobjectives, depending on whether the extracting solutionis directly in contact with the sample or separated by apolymeric membrane which can be held in differentconfigurations. In LPME, furthermore, two-phase andthree-phase systems are possible; the latter can be regardedas a micro liquid-liquid extraction-back-extraction systemwhich exploits the acid-base character of the analytes toachieve simultaneous enrichment and clean-up.

The most representative configurations are presented inFig. 6. More details of their theoretical aspects and generalapplications can be found in recent reviews focussing onthe different modalities of LPME−single drop micro-extraction (SDME) [6], membrane-based extractions [8, 9],and hollow-fibre liquid-phase microextraction (HF-LPME) [5, 7]. Applications of LPME to the extraction ofpolar analytes from water samples are summarized inTables 3 and 4; the applications are discussed below, wheretwo-phase and three-phase systems are treated separately.

Two-phase systems

Applications of two-phase LPME in water analysis arenormally restricted to medium polarity and non-polaranalytes and those whose polarities can be reduced beforeextraction. The reason is simple—the extracting solvent ormixture (acceptor solution) cannot be miscible with water.As a consequence, two-phase LPME is best suited to GCanalysis (Table 3) in contrast with three-phase systemswhich are often employed in combination with LC or CE(Table 4).

The simplest way to perform two-phase LPME is thesingle-drop microextraction mode (SDME). In this ap-

Fig. 5 Dual-phase stir bar for SBSE. Reprinted from Ref. [93],copyright (2005), with permission from Elsevier

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proach, analytes are extracted from the stirred aqueoussample into a drop of organic solvent (ca 1-3 μL)suspended from the needle of a microsyringe (Fig. 6a).After a given time the drop is retracted into the syringe andinjected into the chromatographic system for analysis.Toluene is the solvent most often used, because it is hassome polar character and very low water solubility [96],Table 3. Mixtures of CH2Cl2 and CCl4 have also been

successfully applied to the extraction of organophosphoruscompounds [97]; these solutions are, however, more proneto dissolve or become dislodged when long extractionperiods are used. A final group of solvents with muchpotential in SDME are ionic liquids. These have mainlybeen used for the extraction of PAH, alkylphenoliccompounds, and chloroanilines, before LC determination[98−100]. As discussed in the section on SPME, their

Fig. 6 Different liquid-phase microextraction systems: (a) two-phase single-drop microextraction (2P-SDME), from Ref. [6],copyright (2002), with permission from Elsevier; (b) three-phasesingle-drop microextraction (3P-SDME), from Ref. [114], copyright(2002), with permission from Elsevier; (c) hollow-fibre liquid-phasemicroextraction (HF-LPME) in the rod configuration, from Ref.

[139], copyright (2001), with permission from Elsevier; (d) hollow-fibre liquid-phase microextraction (HF-LPME) in the U-shapeconfiguration, from Ref. [142], copyright (2000) with permissionfrom Wiley-VCH and authors; and (e) flat-membrane microextrac-tion (MMLLE and SLME), from Ref. [119], copyright (2002), withpermission from Elsevier

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Table 3 Applications of two-phase LPME and membrane microextraction to the determination polar species in water samples

Analyte Techniquea E.F.b Acceptor phase Detection Refs.

Nitroaromatic explosives 2P-SDME N.R. Toluene GC-MS [133]2P-HF-LPME N.R. Toluene GC-MS [134]

Organophosphorus and carbamate pesticides 2P-SDME N.R. Toluene and n-hexane LC-UV, GC-MS [102, 135, 136]2P-HF-LPME N.R. Toluene GC-FTD [104]

Organophosphorus warfare agents 2P-SDME N.R. CCl2H2:CCl4 (3:1) GC-MS [97]Antifouling biocides 2P-SDME 33 Toluene GC-ECD [96]Fungicides 2P-HF-LPME 213 Toluene GC-ECD [105, 106]

MMLLE 100 1-Octanol LC-UV [119, 137]Alkylphenols 2P-HF-LPME 163 Toluene GC-MS [108, 138]

2P-HF-LPME 190 1-Octanol GC-MS [138]2P-SDME 163 [C6MIM][PF6] LC-FLD [99]

Phenols 2P-HF-LPME N.R. Ethyl acetate GC-MS [108, 111]2P-SDME 146 Hexyl acetate GC-MS [109]HS-SDME 528 NaOH 1 mol L−1 CE [101]

Acidic pharmaceuticals 2P-HF-LPME 415 1-Octanol GC-MS [138]Aromatic amines HS-SDME N.R. [C4MIM][PF6] LC-UV [100]Triazine herbicides 2P-HF-LPME 208 Toluene GC-MS [103, 110]

2P-SDME 42 Toluene GC-MS [103]Cationic surfactants MMLLE 250 Chlorobutane (10% heptanoic acid) LC-UV [107]

N.R., not reported; [C6MIM][PF6], 1-hexyl-3-methylimidazolium hexafluorophosphate; [C4MIM][PF6], 1-butyl-3-methylimidazoliumhexafluorophosphatea2P, two-phase; HS, headspace; SDME, single-drop microextraction; HF-LPME, hollow-fibre liquid-phase microextraction; MMLLE,microporous membrane liquid-liquid extractionbEnrichment factor: ratio between analyte concentration in the acceptor solution and its initial concentration in the sample. The maximumvalue for the group of analytes considered in each application is reported

Table 4 Applications of three-phase LPME and membrane microextraction to the determination polar species in water samples

Analyte Techniquea E.F.b Organic solventc Detection Refs.

Fungicides SLME 67 DHE with 15% TOPO LC-UV [119, 137]Phenols 3P-HF-LPME 400 1-Octanol LC-UV, MEKC [139–141]

3P-SDME 100 Hexane LC-UV [113, 115]3P-SDME 300 Heptane-toluene (1:1) LC-UV [115]

Acidic pharmaceuticals 3P-HF-LPME 234 1-Octanol LC-MS-MS [112, 122, 142]3P-HF-LPME 15,000d 1-Octanol LC-UV [122]3P-HF-LPME 100 DHE CE [142]

Phenoxy herbicides 3P-HF-LPME 490 1-Octanol LC-UV [143]Glyphosate metabolites SLME 60 DHE doped with methyltrioctylammonium chloride CE [120]Haloacetic acids SLME 60 DHE (5% TOPO) LC-UV [117]

3P-HF-LPME 3,000 DHE doped with DEHP LC-UV [118]Aromatic amines 3P-HF-LPME 6,000d Benzyl alcohol-ethyl acetate (8:2) LC-UV [123, 144]

3P-HF-LPME 500 DHE LC-UV [144]3P-SDME 250 DHE LC-UV [114]3P-SDME 400 Benzyl alcohol-ethyl acetate (2:1) LC-UV [114, 116]

Triazine herbicides SLME 390 DHE LC-UV [145, 146]SLME 140 DHE doped with TOPO LC-UV [146]

Bipyridilium herbicides SLME N.R. DHE doped with DEHP LC-UV [121]

DHE, dihexyl ether; TOPO, trioctyl phosphine oxide; DEHP, di(2-ethylhexyl) phosphoric acida3P, three-phase; HF-LPME, hollow-fibre liquid-phase microextraction; SLME, supported-liquid membrane extractionbEnrichment factor: ratio between analyte concentration in the acceptor solution and its initial concentration in the samplecSolvent impregnating the membrane poresdValues achieved using multiple LPME

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ability to extract more polar analytes may be tuned byselecting the appropriate anions and cations. Ionic liquidshave high viscosity and are virtually non-volatile; stableand larger drops can therefore be used, increasing the yieldof the extraction. These liquids can, moreover, be used inthe headspace mode, with heating of the sample aboveambient temperature [100]. Aqueous solutions of sodiumhydroxide have also be used for the headspace liquidmicroextraction (HS-SDME) of semi-volatile phenols fromacidified water samples; enrichment factors in excess of500 have been achieved for some analytes [101]. Althoughapplications of HS-SDME have been included in Table 3,this mode could, theoretically, also be regarded as a three-phase system in which volatility differences between theanalyte and the other matrix constituents are exploited toachieve a more selective enrichment.

Obviously, some drawbacks of LLE have been inheritedby SDME. One is the possibility of emulsion formationwhen dealing with dirty samples, which in SDME wouldresult in drop dissolution. Drop stability is limited and theprocess requires a dedicated operator, although dynamicautomated systems have been proposed [102]. One way ofminimizing some of these problems is to use a polymericmembrane which serves as a support for the extractingsolvent, enabling the use of larger volumes than in SDME,and a physical barrier between the phases. The membraneis usually made of a porous hydrophobic material(normally polypropylene).

In practice, two approaches can be used to performmembrane-based microextractions. One is the use ofhollow-fibre (HF) membranes, either in a rod like(Fig. 6c) or U-shape (Fig. 6d) configuration; the other isbased on use of a flat membrane (Fig. 6e) to separate thesample (donor) and extracting (acceptor) solutions. Thelatter is normally referred to as microporous membraneliquid-liquid extraction (MMLLE). Both formats have advan-tages and limitations. HF-LPME employs inexpensive mem-branes which are normally discarded after use; automationof the extraction step is difficult, however [5, 7]. InMMLLE applications the membrane is normally reused,with a consequent risk of cross-contamination. This formatof LPME is best suited to on-line hyphenation withchromatographic techniques [8, 9].

Direct comparison of HF-LPME and SDME for deter-mination of triazine herbicides [103] showed that HF-LPME resulted in higher enrichment factors (Table 3) andcleaner extracts and is also less prone to matrix effects[104] than SDME. This is attributed to the membrane porestructure, which reduces the concentration of high-molec-ular-weight compounds in the sample extract.

As with SDME, applications of two-phase HF-LPMEand MMLLE are still limited to medium-polarity com-pounds and those containing ionisable groups, for examplephenols, triazine herbicides, etc. For such compounds,adjustment of sample pH and ionic strength to reduce thesolubility of the analytes are expected to improve the yieldof the extraction. These effects should be carefullyinvestigated for each group of analytes, because reduction

of the kinetics of the extraction at high ionic strengths andhydrolysis of some compounds at extreme pHmight reducethe efficiency of the process [104−106]. An alternative, forextraction of ionic species is to form an ion-pair. Thisstrategy, with heptanoic acid as ion-pairing reagent, hasbeen used to extract cationic surfactants by MMLLE [107].Similar to SPME and SBSE, in-port derivatisation and in-syringe derivatisation can also be combined with LPME[108] and SDME [109], respectively, to improve GCdetermination of some phenolic compounds.

Most applications of membrane LPME have beendeveloped using polymers designed for industrial applica-tions; these have to be cut and, sometimes, sealed in thelaboratory, often resulting in poor reproducibility. As analternative, the company Gerstel (Mülheim, Germany) incollaboration with the Environmental Research Centre(UFZ, Leipzig, Germany) has recently commercialised aset of membranes designed for analytical purposes, whichcan be used in automated operation (MASE). The maindifference between this membrane and the hollow fibresemployed in HF-LPME is that the MASE membranes arenon-porous whereas those normally used in HF-LPME(Accurel Q3/2 from Membrana, Wuppertal, Germany)have a pore size of 0.2 μm. MASE is, therefore, a three-phase system in which the membrane acts as the realinterface. To speed up the mass-transfer kinetics thethickness of the MASE membrane is reduced to 30 μm(compared with 200 μm for that from Accurel) and theextraction temperature is increased to about 40°C [110,111]. Another difference is that MASE membranes aredesigned to accommodate up to 1 mL solvent (comparedwith ca. 10-20 μL in HF-LPME using porous membranes);large-volume injection is, therefore, often needed to reducedetection limits.

Three-phase systems

Three-phase LPME entails extraction of analytes from anaqueous sample to an organic solvent and simultaneousback-extraction from this to the acceptor solution, usually afew microlitres of water at the appropriate pH. The organicsolvent is therefore an interface between both aqueoussolutions. Simultaneous enrichment and clean-up can beachieved by exploiting the acid-base properties of theanalytes, i.e. for acidic analytes the sample is adjusted to apH below the pKa of the compounds to obtain the neutralspecies, which are extracted to the organic solvent, and theacceptor solution to a pH above their pKa, so the analytescan then be converted into the more hydrophilic ionicspecies and trapped in the aqueous extracting solution. Incontrast, if the target species is basic in character, thesample is made basic and the acceptor solution is acidic.The result is enhancement of extraction selectivity, anextremely useful means of eliminating interfering com-pounds when non-selective detectors (for example UV) areemployed and of reducing matrix effects during LC-MSanalysis of complicated environmental samples, for

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example wastewater [112]. As the acceptor solution isaqueous, three-phase LPME is preferably combined withLC and electrodriven separations than to GC. Mostsignificant applications of three-phase LPME for thedetermination of polar species are summarised in Table 4.

As with two-phase LPME, different working modes canbe considered. In three-phase SDME, a layer of organicsolvent is placed between the sample and the acceptorsolution, in direct contact with both. Its volume should beminimised, because it competes with the acceptor solutionfor the analytes. This can be achieved by using a Teflonring (Fig. 6b) [113, 114] or a small volumetric flask [115,116]. This format has been applied to the determination ofphenols [113, 115] and aromatic amines [114, 116]. For thefirst of these non-polar solvents (e.g. hexane, toluene) canbe used whereas amines require more polar solvents (anether, ester, or alcohol). Use of these solvents is, however,more problematic, because they are partially soluble inwater and quite volatile and, as a consequence, if longextraction times are used the enrichment factors obtaineddecrease and precision worsens [114].

Use of porous membranes which contain a trappedorganic solvent to separate the sample and acceptorsolutions seems a more robust configuration for LPME inthree-phase systems. Hollow fibres (three-phase HF-LPME) or flat membranes in flow systems, the latterknown as supported-liquid membrane extraction (SLME),can be used. The set up of both formats is the same as in atwo-phase system (Fig. 6c–e) with the difference that themembrane pores have been previously impregnated withthe organic solvent. The most commonly used solvents are1-octanol and dihexyl ether (DHE), because these arerelatively polar with low water solubility and volatility andform an immobilised organic phase stable for a relativelylong time. For very polar analytes, diffusion through theorganic membrane can be enhanced by doping the organicsolvent with several modifiers. Thus, trioctylphosphineoxide (TOPO), a strong H-bonding agent, has been used topromote the extractability of haloacetic acids [117, 118]and some fungicide metabolites [119]. For extraction ofvery basic/cationic and very acidic/anionic analytes an ion-pair reagent (also named a carrier) can also be employed,for example methyltrioctylammonium chloride for acidic/

Table 5 Comparison of SPE and several microextraction strategies for determination of acidic pharmaceuticals, oestrogens, and phenols inwater samples

Extractiona Determination Sample volume (mL) Processing time (min)b Derivatisationc Matrix effectsd LOD/LOQ (ng L−1) Ref.

Acidic pharmaceuticalsSPE GC-MS 500 90–120 MTBSTFA No 10–50 [147]SPME GC-MS 22 60 On-fibre MTBSTFA Moderate 12–40 [40]2P-LPME GC-MS 5 60 No N.R. 20–40 [138]SPE LC-MS-MS 50 ca 60 No No 0.8–6.5 [148]3P-LPME LC-MS-MS 22 45 No No 0.5–42 [112]OestrogensSPE GC-MS-MS 2,000 ca 180 MSTFA No 1–3 [149]SPME GC-MS-MS 100 90 On-fibre

MSTFAModerateto strong

0.2–3 [39]

SBSE GC-MS-MS 50 240 In-sampleacetylation

No 1–5 [90]

SPE LC-MS-MS 1,000 ca 120 min No No 1–2 [150]IT-SPME LC-MS-MS 1 ca 5 No Moderate 3–12 [76]PhenolsSPE GC-MS 500 60 In-sample

acetylationN.R. 1–10 [151]

SPME GC-MS 12 30 In-sampleacetylation

No 1–61 [63]

SBSE GC-MS 10 120 In-sampleacetylation

No 1–10 [88]

2P-LPME GC-MS 5 30 In-port BSTFA No 5–16 [108]2P-SDME GC-MS 3 20 In-syringe BSA N.R. 4–61 [109]aSPE, solid-phase extraction; SPME, solid-phase microextraction; 2P-LPME, two-phase liquid-phase microextraction; 3P-LPME, three-phase liquid-phase microextraction; SBSE, stir-bar sorptive extraction; IT-SPME, in-tube solid-phase microextraction; 2P-SDME,two-phase single-drop microextractionbApproximate processing time per sample, including evaporation, derivatisation, clean-up, etc., when neededcMTBSTFA, N-methyl-N-(tert-butyldimethylsilyl)trifluoroacetamide; MSTFA, N-methyl-N-(trimethylsilyl)trifluoroacetamide; BSTFA,bis-(trimethylsilyl)trifluoroacetamide; BSA, bis-(trimethylsilyl)acetamidedMatrix effects affecting sample preparation (not determination). N.R., not reported

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anionic analytes [120] or diethylhexyl phosphate forcationic/basic ones [121]. The ion pairs formed areextracted into the organic interface and broken by selectingthe appropriate pH in the acceptor solution, which releasesthe free analytes.

As shown in Table 4, enrichment factors (defined as theratio of the analyte concentrations in the acceptor solutionand in the sample) up to several hundred can be achievedby use of three-phase LPME. For some compounds thesevalues can be improved substantially by performing a two-step LPME [122, 123]. This is achieved by using theaqueous acceptor solution from the first extraction as thedonor phase in the second one, after appropriate pH re-adjustment (acidification for acidic species).

Microextraction strategies comparison and conclusions

As it has been shown throughout this review, the possibilityof using microextraction techniques for analysis of polarorganic contaminants in aqueous samples has beenincreased substantially in recent years. As an example,Table 5 compares some characteristics of different micro-extraction and SPE methods, using GC or LC detection, fordetermination of three groups of polar compounds (acidicpharmaceuticals, oestrogens, and phenols) in water sam-ples. The first point to notice is that microextractiontechniques normally achieve an important decrease notonly in organic waste, but also in sample and timeconsumption. This is very important in the determination ofoestrogens, which normally require extraction of very largesamples and several steps, including clean-up and solventevaporation at several stages when SPE is selected as theconcentration technique.

All the different microextraction techniques availableseem to result in similar detection limits. SPME and SDMEare the most sensitive techniques toward matrix effects.

Although derivatisation of polar analytes can beperformed in combination with most microextractionstrategies, possibilities are greater with SPME and, there-fore, this is often the technique of choice if the analytes areto be detected by GC. Although use of derivatisationreactions improves the extractability of some analytes, thedevelopment of new coatings is expected to enhance theapplicability of SPME to the determination of polarspecies. Sol-gel polymerisation seems to be the mostpromising method for immobilisation of polar sorbents onmicroextraction fibres. The coated phases obtained can berinsed with polar solvents and are more stable than those oncommercial fibres. They are, therefore, suitable for use incombination with both GC and LC. Lack of alternativecoatings to PDMS is clearly limiting the applicability ofSBSE to polar compounds.

If analytes are to be detected by LC or CE, the mostpromising technique seems to be three-phase LPME usingporous membranes impregnated with the appropriatesolvent. This enables simultaneous enrichment and clean-up, important not only for non-selective detection (e.g. UV)but also for MS detection, because it may help to minimise

the matrix effects that complicate quantification by LC-MS. Currently, however, its main shortcoming is the use oflaboratory-designed extraction devices, which requireskilled operators, increase the variability of the results,and slow the spread of the technique outside researchlaboratories. It is believed that commercialisation of thetechnique, in a limited number of formats, will overcomesuch problems.

Acknowledgements Financial support by the Spanish DGICTMinisterio de Educación y Ciencia (project BQU 2003-02090) andXunta de Galicia (project PGIDIT03TAM02E) is acknowledged.J.B. Quintana is grateful toMinisterio de Educación y Ciencia for hispostdoctoral grant.

References

1. Schmidt CK, Fleig M, Sacher F, Brauch HE (2004) EnvironPollut 131:107–124

2. Pawliszyn J (1997) Solid phase microextraction: theory andpractice. Wiley-VCH, New York

3. Pawliszyn J (2002) In: Pawliszyn J (ed) Solid phasemicroextraction. Elsevier, Amsterdam, pp 389–477

4. Baltussen E, Sandra P, David F, Cramers C (1999) JMicrocolumnSep 11:737–747

5. Psillakis E, Kalogerakis N (2003) Trends Anal Chem 22:565–5746. Psillakis E, Kalogerakis N (2002) Trends Anal Chem 21:53–637. Rasmussen KE, Pedersen-Biergaard S (2004) Trends Anal

Chem 23:1–108. Jönsson JA, Mathiasson L (2001) J Sep Sci 24:495–5079. Jönsson JA, Mathiasson L (2003) LC GC Eur 16:683–69010. Prosen H, Zupanèiè-Kralj L (1999) Trends Anal Chem

18:272–28211. Fontanals N, Marcé RM, Borrull F (2005) Trends Anal Chem

24:394–40612. Chong SL, Wang DX, Hayes JD, Wilhite BW, Malik A (1997)

Anal Chem 69:3889–389813. Kim TY, Alhooshani K, Kabir A, Fries DP, Malik A (2004)

J Chromatogr A 1047:165–17414. Alhooshani K, Kim TY, Kabir A, Malik A (2005) J Chromatogr

A 1062:1–1415. Bigham S, Medlar J, Kabir A, Shende C, Alli A, Malik A

(2002) Anal Chem 74:752–76116. Liu MM, Zeng ZR, Wang CL, Tan YJ, Liu H (2003)

Chromatographia 58:597–60517. Zuin VG, Lopes AL, Yariwake JH, Augusto F (2004)

J Chromatogr A 1056:21–2618. Kabir A, Hamlet C, Malik A (2004) J Chromatogr A 1047:1–1319. Yun L (2003) Anal Chim Acta 486:63–7220. Wang DH, Xing J, Peng JG, Wu CY (2003) J Chromatogr A

1005:1–1221. Yu JX, Wu CY, Xing J (2004) J Chromatogr A 1036:101–11122. Cai LS, Zhao YY, Gong SL, Dong L, Wu CY (2003)

Chromatographia 58:615–62123. Li XJ, Zeng ZR, Gao SZ, Li HB (2004) J Chromatogr A

1023:15–2524. Wang W, Gong SL, Cao QH, Chen YY, Li XJ, Zeng ZR (2005)

Chromatographia 61:75–8025. Liu M, Zeng Z, Fang H (2005) J Chromatogr A 1076:16–2626. Liu MM, Zeng ZR, Tian Y (2005) Anal Chim Acta 540:341–35327. Basheer C, Jegadesan S, Valiyaveettil S, Lee HK (2005)

J Chromatogr A 1087:252–25828. Djozan D, Assadi Y, Karim-Nezhad G (2002) Chromatographia

56:611–61629. Farajzadeh MA, Rahmani NA (2005) Talanta 65:700–70430. Wu JC, Lord HL, Pawliszyn J, Kataoka H (2000) J Microcolumn

Sep 12:255–266

1459

31. Wu JC, Pawliszyn J (2001) J Chromatogr A 909:37–5232. Wu JC, Pawliszyn J (2004) Anal Chim Acta 520:257–26433. Bagheri H, Mir A, Babanezhad E (2005) Anal Chim Acta

532:89–9534. Ghassempour A, Najafi NM, Mehdinia A, Davarani SSH,

Fallahi M, Nakhshab M (2005) J Chromatogr A 1078:120–12735. Liu JF, Li N, Jiang GB, Li JM, Jönsson JA, Wen MJ (2005)

J Chromatogr A 1066:27–3236. Anderson JL, Armstrong DW (2003) Anal Chem 75:4851–

485837. Braun P, Moeder M, Schrader S, Popp P, Kuschk R, Engewald

W (2003) J Chromatogr A 988:41–5138. Moeder M, Schrader S, Winkler M, Popp P (2000) J Chromatogr

A 873:95–10639. Carpinteiro J, Quintana JB, Rodriguez I, Carro AM, Lorenzo

RA, Cela R (2004) J Chromatogr A 1056:179–18540. Rodríguez I, Carpinteiro J, Quintana JB, Carro AM, Lorenzo

RA, Cela R (2004) J Chromatogr A 1024:1–841. Krutz LJ, Senseman SA, Sciumbato AS (2003) J Chromatogr A

999:103–12142. Stashenko EE, Martínez JR (2004) Trends Anal Chem 23:

553–56143. Bao ML, Pantani F, Griffini O, Burrini D, Santianni D, Barbieri

K (1998) J Chromatogr A 809:75–8744. Cancho B, Ventura F, Galceran MT (2002) J Chromatogr A

943:1–1345. Tsai SW, Chang CM (2003) J Chromatogr A 1015:143–15046. Salgado-Petinal C, Alzaga R, García-Jares C, Llompart M,

Bayona JM (2005) Int J Environ Anal Chem 85:543–55247. Salgado-Petinal C, Alzaga R, García-Jares C, Llompart M,

Bayona JM (2005) Anal Chem 77:6012–601848. Wittmann G, Van Langenhove H, Dewulf J (2000) J Chromatogr

A 874:225–23449. Pan L, Pawliszyn J (1997) Anal Chem 69:196–20550. Aikawa B, Burk RC (1997) Int J Environ Anal Chem 66:215–22451. Sarrión MN, Santos FJ, Galceran MT (1999) J Chromatogr A

859:159–17152. Sarrión MN, Santos FJ, Galceran MT (2000) Anal Chem

72:4865–487353. Nilsson T, Baglio D, Galdo-Miguez I, Madsen JO, Facchetti S

(1998) J Chromatogr A 826:211–21654. Henriksen T, Svensmark B, Lindhardt B, Juhler RK (2001)

Chemosphere 44:1531–153955. Lee MR, Lee RJ, Lin YW, Chen CM, Hwang BH (1998) Anal

Chem 70:1963–196856. Alzaga R, Peña A, Bayona JM (2003) J Sep Sci 26:87–9657. Sng MT, Ng WF (1999) J Chromatogr A 832:173–18258. Rodríguez I, Rubí E, González R, Quintana JB, Cela R (2005)

Anal Chim Acta 537:259–26659. Alzaga R, Peña A, Ortiz L, Bayona JM (2003) J Chromatogr A

999:51–6060. Alzaga R, Bayona JM (2004) J Chromatogr A 1042:155–16261. Canosa P, Rodríguez I, Rubi E, Cela R (2005) J Chromatogr A

1072:107–11562. Buchholz KD, Pawliszyn J (1994) Anal Chem 66:160–16763. Llompart M, Lourido M, Landín P, García-Jares C, Cela R

(2002) J Chromatogr A 963:137–14864. Okeyo PD, Snow NH (1998) J Microcolumn Sep 10:551–55665. Lamas JP, Salgado-Petinal C, García-Jares C, Llompart M, Cela

R, Gómez M (2004) J Chromatogr A 1046:241–24766. Pan L, Chong JM, Pawliszyn J (1997) J Chromatogr A

773:249–26067. Zhao YY, Cai LS, Jing ZZ, Wang H, Yu HX, Zhang HS (2003)

J Chromatogr A 1021:175–18168. Zimmermann T, Ensinger WJ, Schmidt TC (2004) Anal Chem

76:1028–103869. Huang MK, Liu CR, Huang SD (2002) Analyst 127:1203–120670. Chia KJ, Huang SD (2005) Anal Chim Acta 539:49–5471. Saito Y, Jinno K (2003) J Chromatogr A 1000:53–6772. Zambonin CG (2003) Anal Bioanal Chem 375:73–8073. Popp P, Bauer C, Möder M, Paschke A (2000) J Chromatogr A

897:153–159

74. Gou YN, Eisert R, Pawliszyn J (2000) J Chromatogr A873:137–147

75. Kataoka H (2002) Anal Bioanal Chem 373:31–4576. Mitami K, Fujioka M, Kataoka H (2005) J Chromatogr A

1081:218–22477. Reemtsma T (2001) Trends Anal Chem 20:533–54278. Shintani Y, Zhou X, Furuno M, Minakuchi H, Nakanishi K

(2003) J Chromatogr A 985:351–35779. Tan BCD, Marriott PJ, Lee HK, Morrison PD (1999) Analyst

124:651–65580. Globig D, Weickhardt C (2005) Anal Bioanal Chem 381:656–65981. Standler A, Schatzl A, Klampfl CW, Buchberger W (2004)

Microchim Acta 148:151–15682. Nakamura S, Daishima S (2005) Anal Bioanal Chem 382:99–10783. Serôdio P, Nogueira JMF (2004) Anal Chim Acta 517:21–3284. Itoh N, Tao H, Ibusuki T (2005) Anal Chim Acta 535:243–25085. Kawaguchi M, Inoue K, Yoshimura M, Ito R, Sakui N,

Okanouchi N, Nakazawa H (2004) J Chromatogr B 805:41–4886. Nakamura S, Daishima S (2004) J Chromatogr A 1038:291–29487. Kawaguchi M, Inoue K, Yoshimura M, Sakui N, Okanouchi N,

Ito R, Yoshimura Y, Nakazawa H (2004) J Chromatogr A1041:19–26

88. Kawaguchi M, Ishii Y, Sakui N, Okanouchi N, Ito R, Saito K,Nakazawa H (2005) Anal Chim Acta 533:57–65

89. Montero L, Conradi S, Weiss H, Popp P (2005) J ChromatogrA 1071:163–169

90. Kawaguchi M, Ishii Y, Sakui N, Okanouchi N, Ito R, Inoue K,Saito K, Nakazawa H (2004) J Chromatogr A 1049:1–8

91. Kawaguchi M, Sakui N, Okanouchi N, Ito R, Saito K,Nakazawa H (2005) J Chromatogr A 1062:23–29

92. Kawaguchi M, Ito R, Sakuai N, Okanouchi N, Saito K,Nakazawa H (2006) J Chromatogr A 1105:140–147

93. Bicchi C, Cordero C, Liberto E, Rubiolo P, Sgorbini B, DavidF, Sandra P (2005) J Chromatogr A 1094:9–16

94. Montero L, Popp P, Paschke A, Pawliszyn J (2004) J ChromatogrA 1025:17–26

95. Popp P, Bauer C, Paschke A, Montero L (2004) Anal Chim Acta504:307–312

96. Lambropoulou DA, Albanis TA (2004) J Chromatogr A 1049:17–23

97. Palit M, Pardasani D, Gupta AK, Dubey DK (2005) Anal Chem77:711–717

98. Liu J, Jiang GB, Chi YG, Cai YQ, Zhou QX, Hu JT (2003) AnalChem 75:5870–5876

99. Liu JF, Chi YG, Jiang GB, Tai C, Peng JF, Hu JT(2004) J Chromatogr A 1026:143–147

100. Peng JF, Liu JF, Jiang GB, Tai C, Huang MJ (2005) J ChromatogrA 1072:3–6

101. Zhang J, Su T, Lee HK (2005) Anal Chem 77:1988–1992102. Guo L, Liang P, Zhang TZ, Liu Y, Liu S (2005)

Chromatographia 61:523–526103. Shen G, Lee HK (2002) Anal Chem 74:648–654104. Lambropoulou DA, Albanis TA (2005) J Chromatogr A

1072:55–61105. Pan HJ, Ho WH (2004) Anal Chim Acta 527:61–67106. Lambropoulou DA, Albanis TA (2004) J Chromatogr A

1061:11–18107. Norberg J, Thordarson E, Mathiasson L, Jönsson JA (2000)

J Chromatogr A 869:523–529108. Basheer C, Lee HK (2004) J Chromatogr A 1057:163–169109. Saraji M, Bakhshi M (2005) J Chromatogr A 1098:30–36110. Hauser B, Popp P, Kleine-Benne E (2002) J Chromatogr A

963:27–36111. Schellin M, Popp P (2005) J Chromatogr A 1072:37–43112. Quintana JB, Rodil R, Reemtsma T (2004) J Chromatogr A

1061:19–26113. Zhao LM, Lee HK (2001) J Chromatogr A 931:95–105114. Zhu LY, Tay CB, Lee HK (2002) J Chromatogr A 963:231–237115. Sarafraz-Yazdi A, Beiknejad D, Es’haghi Z (2005)

Chromatographia 62:49–54116. Sarafraz Yazdi A, Es’haghi Z (2005) Talanta 66:664–669

1460

117. Wang XY, Saridara C, Mitra S (2005) Anal Chim Acta 543:92–98

118. Kou DW, Wang XY, Mitra S (2004) J Chromatogr A 1055:63–69119. Sandahl M, Mathiasson L, Jönsson JA (2002) J Chromatogr A

975:211–217120. Dżygiel P, Wieczorek P (2001) J Sep Sci 24:561–566121. Mulugeta M, Megersa N (2004) Talanta 64:101–108122. Wen XJ, Tu CH, Lee HK (2004) Anal Chem 76:228–232123. Sarafraz Yazdi A, Es’haghi Z (2005) J Chromatogr A

1082:136–142124. Pan L, Adams M, Pawliszyn J (1995) Anal Chem 67:4396–

4403125. Díaz A, Ventura F, Galceran MT (2002) Anal Chem 74:3869–

3876126. Eisert R, Pawliszyn J (1997) Anal Chem 69:3140–3147127. Wu JC, Tragas C, Lord H, Pawliszyn J (2002) J Chromatogr A

976:357–367128. Takino M, Daishima S, Nakahara T (2001) Analyst 126:602–608129. Gou YN, Tragas C, Lord H, Pawliszyn J (2000) J Microcolumn

Sep 12:125–134130. Gou YN, Pawliszyn J (2000) Anal Chem 72:2774–2779131. Wu JC, Pawliszyn J (2001) Anal Chem 73:55–63132. Wu JC, Mester Z, Pawliszyn J (2000) Anal Chim Acta

424:211–222133. Psillakis E, Kalogerakis N (2001) J Chromatogr A 938:113–120134. Psillakis E, Mantzavinos D, Kalogerakis N (2004) Anal Chim

Acta 501:3–10

135. Liang P, Guo L, Liu Y, Liu S, Zhang TZ (2005) Microchem J80:19–23

136. Lambropoulou DA, Psillakis E, Albanis TA, Kalogerakis N(2004) Anal Chim Acta 516:205–211

137. Sandahl M, Mathiasson L, Jönsson JA (2000) J Chromatogr A893:123–131

138. Müller S, Möder M, Schrader S, Popp P (2003) J ChromatogrA 985:99–106

139. Zhu LY, Zhu L, Lee HK (2001) J Chromatogr A 924:407–414140. Zhu LY, Tu CH, Lee HK (2001) Anal Chem 73:5655–5660141. Jiang XM, Oh SY, Lee HK (2005) Anal Chem 77:1689–1695142. Pedersen-Bjergaard S, Rasmussen KE (2000) Electrophoresis

21: 579–585143. Wu JM, Ee KH, Lee HK (2005) J Chromatogr A 1082:121–127144. Zhao LM, Zhu LY, Lee HK (2002) J Chromatogr A 963:239–248145. Megersa N, Solomon T, Jönsson JA (1999) J Chromatogr A

830:203–210146. Megersa N, Chimuka L, Solomon T, Jönsson JA (2001) J Sep

Sci 24:567–576147. Rodríguez I, Quintana JB, Carpinteiro J, Carro AM, Lorenzo

RA, Cela R (2003) J Chromatogr A 985:265–274148. Quintana JB, Reemtsma T (2004) Rapid Commun Mass

Spectrom 18:765–774149. Quintana JB, Carpinteiro J, Rodriguez I, Lorenzo RA, Carro

AM, Cela R (2004) J Chromatogr A 1024:177–185150. Ingrand V, Herry G, Beausse J, de Roubin MR (2003)

J Chromatogr A 1020:99–104151. Schlett C, Pfeifer B (1992) Vom Wasser 79:65–74

1461