sample handling strategies for the determination of persistent trace organic contaminants from biota...

A

sblcss(as©

K

C

0d

Analytica Chimica Acta 590 (2007) 1–16

Review

Sample handling strategies for the determination of persistenttrace organic contaminants from biota samples

Natalia Fidalgo-Used, Elisa Blanco-Gonzalez, Alfredo Sanz-Medel ∗Department of Physical and Analytical Chemistry, University of Oviedo, C/Julian Claverıa 8, 33006 Oviedo, Spain

Received 28 November 2006; received in revised form 28 February 2007; accepted 2 March 2007Available online 12 March 2007

bstract

Even after emergence of most advanced instrumental techniques for the final separation, detection, identification and determination of analytes,ample handling continues to play a basic role in environmental analysis of complex matrices. In fact, sample preparation steps are often theottleneck for combined time and efficiency in many overall analytical procedures. Thus, it is not surprising that, in the last two decades, aot of effort has been devoted to the development of faster, safer, and more environment friendly techniques for sample extraction and extractlean up, prior to actual instrumental analysis. This article focuses on the state of the art in sample preparation of environmental solid biologicalamples dedicated to persistent organic pollutants (POPs) analysis. Extraction techniques such as Soxhlet extraction, sonication-assisted extraction,
upercritical fluid extraction (SFE), microwave-assisted extraction (MAE), pressurised liquid extraction (PLE) and matrix solid-phase dispersionMSPD) are reviewed and their most recent applications to the determination of POPs in biota samples are provided. Additionally, classical as wells promising novel extraction/clean-up techniques such as solid phase microextraction (SPME) are also summarized. Finally, emerging trends inample preparation able to integrate analytes extraction and their adequate clean-up are presented.
2007 Elsevier B.V. All rights reserved.

eywords: Biota samples; Sample preparation; Sample handling; Extract clean-up; Extraction; Persistent organic pollutants

ontents

1. Introduction. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22. Sample-extraction techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2

2.1. Conventional Soxhlet extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2. Sonication-assisted extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.3. Supercritical fluid extraction (SFE). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42.4. Microwave-assisted extraction (MAE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52.5. Pressurised liquid extraction (PLE). . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62.6. Matrix solid-phase dispersion (MSPD) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7

3. Extract clean-up . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83.1. Classical liquid adsorption chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83.2. Gel-permeation chromatography (GPC) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113.3. Solid-phase microextraction (SPME) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

4. Integrated extraction and clean-up . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135. Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14

∗ Corresponding author. Tel.: +34 98 5103474; fax: +34 98 5103125.E-mail address: [email protected] (A. Sanz-Medel).

003-2670/$ – see front matter © 2007 Elsevier B.V. All rights reserved.oi:10.1016/j.aca.2007.03.004

mailto:[email protected]

dx.doi.org/10.1016/j.aca.2007.03.004

2 tica C

1

g(dacanobosctprttPcmb(sctgbrc

c(osqstc[cmcssecottp

opt

dfmTopa

cfio1tmpcs

ddoat

2

aatpls

seaala

2

dtasarnps

N. Fidalgo-Used et al. / Analy

. Introduction

Persistent organic pollutants (POPs) are a heterogeneousroup of substances including, polychlorinated biphenylsPCBs), polychlorinated dibenzo-p-dioxins and polychlorinatedibenzofurans (PCDD/Fs), organochlorine pesticides (OCPs)nd other organic pollutants such as polycyclic aromatic hydro-arbons (PAHs) [1,2]. POPs toxicity has been linked to observeddverse effects on human health and animals, including cancer,ervous system damage, reproductive disorders and disruptionf the immune system [3,4]. Moreover, they are characterizedy a high chemical and biological stability, and a high degreef lipophilicity. These characteristics make POPs prone to per-ist in the environment and to bioaccumulate along the foodhain involving a wide range of trophic levels [5]. Becausehey circulate globally via the atmosphere, oceans, and otherathways, POPs released in one part of the world can travel toegions far from their source of origin [1,2]. Therefore, despitehe intensive research carried out worldwide during the lasthree decades, it is still needed to improve our knowledge onOPs contamination, since released POPs will continue to cir-ulate in our environment for many years to come. In addition,any of more recently recognized pollutants, such as poly-

rominated diphenylethers (PBDEs), polybrominated biphenylsPBBs), polychlorinated naphthalenes (PCNs), perfluorooctaneulfonate (PFOS) and related compounds, nitro musks and poly-yclic musks fragrances, have distribution patterns similar tohose of conventional POPs mentioned above with regard tolobal biospheric distribution, persistence, bioaccumulation andiomagnification [6]. Consequently, concern about the envi-onmental fate and potential effects of these emerging POPsontinues to increase [7,8].

The analytical procedures used for the determination of bothonventional and emerging POPs in environmental sampleswater, soil, sediment and biota) typically involve a numberf equally relevant steps for sampling, sample preparation,eparation and detection of target compounds, identification,uantification and data handling. The separation and detectiontep has been dominated by gas chromatography (GC) coupledo sensitive and specific detection systems, such as the electronapture detector (ECD) and mass spectrometry detection (MS)9]. Nowadays, these chromatographic instrumental techniquesan provide high resolution of complex mixtures on almost everyatrix, while detection limits below femtograms of the desired

ompound are achievable. Notwithstanding the importance ofuch instrumental techniques, the whole analytical process cantill be wasted if an unsuitable sample preparation procedure ismployed before the sample reaches the detector. This is espe-ially important in environmental analysis where the complexityf many matrices, e.g. biota, and the low concentrations at whichhe analytes of interest have to be identified and/or quantified (inhe presence of many other closely related compounds) makerior sample treatment mandatory.

Sample preparation is usually a multi-step procedure carriedut off-line. This fact makes it tedious and time-consuming,rone to loss of analytes and to contamination. Therefore,oday sample preparation is still the weakest link and the time-

am

p

himica Acta 590 (2007) 1–16

etermining step in the whole analytical procedure, accountingor about two-thirds of the total analysis time. It is also the pri-ary source of errors and discrepancies between laboratories.hus, the quality of this step is a key factor in the final successf the analysis and the judicious choice of an appropriate sam-le preparation procedure greatly influences the reliability andccuracy of a given environmental analysis [10,11].

The basic concept of a sample preparation is to convert aomplex matrix into a sample in a format that is suitable fornal instrumental analysis. This can be achieved by a wide rangef techniques, many of which have changed little over the last00 years [12]. All those sample preparation techniques havehe following common aims: to extract the analytes from the

atrix, to bring them to a suitable concentration level, to removeossible interferences (clean-up step) and, when is required, toonvert the analytes into a more suitable form for detection oreparation.

This paper reviews the main extraction and clean-up proce-ures, published in the last 5 years, applied to the analyticaletermination of both conventional and emerging persistentrganic pollutants (POPs) in environmental biota (vegetal andnimal) samples. Readers interested in older revisions on theopic are advised to consult previous reviews [13–15].

. Sample-extraction techniques

The analysis of POPs in biota samples requires the use ofnalyte extraction techniques, which allow the release of thenalytes from the solid matrix, with optimum yield and selec-ivity, by using an adequate solvent, in such a way that as fewotential interfering species as possible are carried into the ana-ytical separation stage. The solvents may be organic liquids,upercritical fluids, pressurised, or superheated liquids.

The most common extraction techniques for solid and semi-olid matrices include Soxhlet extraction, sonication-assistedxtraction, supercritical fluid extraction (SFE), microwave-ssisted extraction (MAE), pressurized liquid extraction (PLE)nd matrix solid-phase dispersion (MSPD). Thus, we will reviseately reported strategies and applications, using each of suchpproaches.

.1. Conventional Soxhlet extraction

Soxhlet extraction is a general and well-established techniqueeveloped in 1879. The technique is based on exhaustive extrac-ion of organic compounds (analytes) in a Soxhlet system byn organic solvent, which is continuously refluxed through theample contained in a porous thimble. The extracted analytesccumulate in a heated flask and so they must be stable in theefluxing boiling solvent. Soxhlet extraction is the oldest tech-ique used for the isolation of non-polar and semi-polar organicollutants from different types of solid matrices, including biotaamples [10,13,16]. It has been used for decades and has been
dopted by the U.S. Environmental Protection Agency (EPA) asethod 3540C [2].Although the size of the system can vary, the more common
rocedures use 50–200 mL of organic solvent to extract the ana-

tica C

leataoasrtmsvf

urdiwaltStdpebMseaoam

wdut(faavutO(afia(afe

Sfihh[faritaefPP

2

tsupsf2rcovuumoeataepdu

muud(elpdl


ytes from between 1 and 100 g of biological tissue [10]. It isssential to match the solvent polarity to the solute solubilitynd to thoroughly wet the sample matrix with the solvent forhe extraction. Typical solvents to extract POPs from animalnd plant tissues are n-hexane, dichloromethane, and mixturesf toluene–methanol, n–hexane–acetone and dichloromethane-cetone. Usually, animal and plant fresh tissues should be cut,hredded and then ground with sodium sulphate in order toeduce their water content and helps to open up the tissue struc-ure which enables good penetration of solvent into the sample

atrix [10]. As alternative to chemical drying with sodiumulphate, freeze-drying (water evaporation below 0 ◦C underacuum conditions) or liophylization of samples can be per-ormed before extraction.

The advantages of Soxhlet extraction include: it allows these of large amount of sample (e.g. 1–100 g), no filtration isequired after the extraction, the technique is not matrix depen-ent, and many Soxhlet extractors can be set up to performn unattended operation. Attempts to automate the techniqueere somewhat successful, and a few commercial systems are

vailable in which several samples can be extracted in paral-el with much shorter extraction times and less organic solventhan using conventional Soxhlet [16]. The main disadvantages ofoxhlet extraction are that it requires large amounts of solvent,

he solvent must be evaporated to concentrate analytes beforeetermination, the process takes several hours or days to com-lete the extraction, and it generates dirty extracts that requirextensive clean-up [13,16]. Therefore, this traditional method iseing replaced by other new extraction techniques such as SFE,AE and ASE with shortened extraction times, reduced organic

olvent consumption and increased pollution prevention. How-ver, Soxhlet extraction is still an attractive option for routinenalysis for its general robustness and relatively low cost. More-ver, Soxhlet extraction is widely used as a standard techniquend reference for evaluating the performance of new extractionethods proposed.Soxhlet extraction was used recently to extract PAHs from

hole gall bladders and liver of different fish species usingichloromethane as solvent [17]. Dichloromethane was alsosed to extract OCPs and PCBs from fish and bowhead whaleissue [18], OCPs, PCBs and PBDEs from several biota samplesmussel, fish, birds, mammals, lichen) [19], PCBs and PCDD/Fsrom albatross muscle [20] and PBDEs from salmon [21]. Vivesnd Grimalt [22] reported the Soxhlet extraction of PAHs, PCBsnd OCPs from fish liver with n-hexane–dichloromethane (4:1,/v). Different n-hexane–dichloromethane mixtures were alsosed for extracting OCPs and PCBs from krill and silverfishissue [23], non-ortho PCBs congeners from fish tissue [24],CPs, PCBs, PCDD/Fs and PCNs from several animal species

polar bear, krill, sharp-spined notothen) [25], PCBs, PCDD/Fsnd PCNs from cormorants eggs and gulls [26], PBDEs fromsh tissue [27,28], from fish muscle, fish liver and mussel [29]nd polycyclic musk fragances from marine mammals tissue
bear, seal, sea lion, sea otters, dolphins) [30]. Manirakiza etl. [31] used a Soxhlet technique to extract OCPs and PCBsrom seven fish species with n-hexane–acetone (3:1, v/v). Sev-ral n-hexane–acetone mixtures also allowed the extraction by
c5da

himica Acta 590 (2007) 1–16 3

oxhlet of PBDEs from biota [32,33], from shellfish tissue,sh muscle and fish liver [34] and from cormorant liver andarbour porpoises [35]. Similarly, PBDEs, PCBs and OCPsave been extracted from guillemot liver and guillemot eggs36], PCBs from mullet fish [37] and coots tissue [38], OCPsrom salmon trout [39] and brominated flame retardants suchs hexabromocyclododecane (HBCD) from biota [40]. Mostecently, Holmqvist et al. [41] reported the use of n-hexanen a Soxhlet apparatus to extract PCBs and DDTs from eelissue while Ramu et al. [42] extracted PBDEs from dolphinnd porpoise tissues (blubber, liver and kidney) with diethylther–n-hexane. Soxhlet was used to extract PCBs and PCDD/Fsrom herring [43], PCBs and PCNs from pine needles [44],CBs, PBDEs and PCNs from mussel [45] and PCBs, PCDD/Fs,BDEs and PCNs from fish [46] using toluene.

.2. Sonication-assisted extraction

The simplest solid–liquid extraction technique is to blendhe solid sample with an appropriate organic solvent and ultra-onicate them. The process is carried out in discrete systemssing an ultrasonic bath or a closed extractor fitted with a sonicrobe. Sonication involves the use of sound waves to stir theample immersed in the organic solvent. Briefly, energy in theorm of acoustic sound waves in the ultrasound region above0 kHz, is used to accelerate mass transport and mechanicalemoval of analytes from the solid matrix surface by a processalled “cavitation”. This consists of the formation and implosionf vacuum bubbles trough the solvent, thus creating microen-ironments with high temperatures and pressures (estimatedp to 5000 ◦C and 100 MPa) [47]. This mechanical effect ofltrasound induces a greater penetration of solvent into solidaterials and improves mass transfer leading to an enhancement

f sample extraction efficiency. Therefore, sonication-assistedxtraction is faster (5–30 min for sample) than the Soxhlet modend allows extraction of large amounts of sample with a rela-ively low cost. Unfortunately, it still uses about as much solvents the Soxhlet extraction, and also filtration is required afterxtraction. Moreover, it is labour intensive since, apart from theolarity of the solvent, the efficiency of the extraction is depen-ent upon the nature and homogeneity of the sample matrix, theltrasound frequency and the sonication time used [13].

Sonication-assisted extraction has been approved by EPA asethod 3550B [2]. Rodrıguez-Sanmartın et al. [48] reported

ltrasound-assisted extraction of PAHs from mussel soft tissuesing dichloromethane. PAHs were also extracted from pine nee-les by ultrasonic extraction with n-hexane–dichloromethane1:1, v/v) [49]. The procedure was compared with Soxhletxtraction and PLE using the same solvent. Despite similar ana-ytical results obtained, the ultrasonic procedure was said torevail over the other two tested in terms of availability, expe-itiousness for simultaneous extractions of various samples andess instrumental cost. A fast ultrasound-assisted extraction pro-
edure, using a mixture of methanol–water (4:1, v/v) containing% triethylamine (TEA) as extraction solvent, was used for theetermination of 14 chlorophenols in clam tissue [50]. Smith etl. [51] studied the influence of the extraction methodology on

4 tica Chimica Acta 590 (2007) 1–16

tepTb

uuTlcawisioaauvtm

tswp[

2

cbfimstmph

ttpoststflmcflrs

F[

tr7flAm

atsted(csmdi

nrSesfpcSfaepon4C


he determination of PAHs in pasture vegetation. The extractionfficiencies of sonication and Soxhlet procedures were com-ared again here using dichloromethane as extraction solvent.he PAHs total amounts extracted by sonication turned out toe only 22–50% of those accessible by Soxhlet.

Ultrasonic-assisted extraction has been recently carried outsing a dynamic extraction set-up (a flow system) which contin-ously supplies fresh extraction solvent to the extraction vessel.his approach may be considered as if it forces adsorbed ana-

ytes to partition continuously into new extraction solvent. Aonsiderable reduction of extraction time, solvent consumptionnd sample handling, with respect to the extraction in static way,as reported [47]. Another feature of such dynamic arrangement

s that the analytes are transferred out of the extraction vesselystem as soon as they are extracted. This can be especiallymportant to avoid degradation of the analytes due to sonicationr if thermo-labile analytes are extracted at higher temperaturesnd pressures. Domeno et al. [52] used a dynamic sonication-ssisted extraction procedure for extracting PAHs from lichenssing hexane. The reported total extraction time was only 10 minersus 2 h in the static extraction mode and 6 h in Soxhlet extrac-ion, while the PAHs relative recovery obtained by the three

ethods was similar.Similar to Soxhlet technique, only drying and homogeniza-

ion is carried out before sonication-assisted extraction of biotaamples. Drying the samples is performed by evaporation ofater at room temperature [52] or by grinding with sodium sul-hate [49]. Freeze-drying can be also used for sample drying48,50].

.3. Supercritical fluid extraction (SFE)

SFE is an extraction technique that uses a solvent in its super-ritical state. Supercritical fluids have similar densities to liquids,ut lower viscosities and so analytes show higher diffusion coef-cients. This combination of properties results in a fluid that isore penetrating has a higher solvating power and may extract

olutes faster and more efficiently than liquids [12–14]. In addi-ion, the density (and therefore the solvent power of the fluid)

ay be adjusted by varying both the pressure and the tem-erature, affording the opportunity of theoretically performingighly selective extractions [14].

SFE utilises commercially available equipments [14] wherehe fluid is pumped, at a pressure above its critical point, throughhe sample placed in an inert extraction cell (see Fig. 1). The tem-erature of the cell is increased to overcome the critical valuef the fluid. After depressurization, analytes are collected in amall volume of organic solvent or on a solid-phase filled car-ridge (solid adsorbent trap). Extraction can be performed intatic, dynamic or recirculating mode: performing static extrac-ion the cell containing the sample is filled with the supercriticaluid, pressurised and allowed to equilibrate; using a dynamicode, the supercritical fluid is passed through the extraction

ell continuously; finally in the recirculating mode the sameuid is repeatedly pumped through the sample and, after theequired number of cycles, it is pumped out to the collectionystem.

epte

ig. 1. Schematic diagram of a SFE system. Reprinted with permission from22].

Although many supercritical fluids have been investigated,he most commonly used is carbon dioxide (CO2) because iteaches the supercritical state at a relatively low pressure (i.e.MPa) and temperature (i.e. 31.3 ◦C), it is non-toxic, non-ammable, non-corrosive, chemically very inert, and affordable.lso, although CO2 is non-polar, its polarity can be adjusted withodifiers such as acetone and methanol.SFE efficiency is affected by a wide range of parameters such

s supercritical fluid nature, temperature and pressure, extrac-ion time, the shape of the extraction cell, the sample particleize, the matrix type, the moisture content of the matrix andhe analyte collection system. Due to these numerous param-ters affecting the extraction efficiencies, SFE affords a highegree of selectivity and the extracts are relatively quite cleanthus, they require only moderate additional clean-up). In fact,ombined with solid adsorbent traps, SFE may provide a single-tep extraction and clean-up. However, the need to control soany operating parameters makes SFE optimization tedious and

ifficult in practice. Other disadvantages of the SFE techniquenclude: limited sample size and high cost of the equipment.

Since, the early use of SFE for extraction in the mid-1980s,umerous applications of the technique in the analysis of envi-onmental samples has been reported [12,13,14]. In addition,FE has been adopted by the EPA as a reference method forxtracting PAHs (Method 3561) and PCBs (Method 3562) fromolid environmental matrices [2]. SFE was also used recentlyor extracting POPs from different plant materials [53–57]. Thelant samples were air dried at room temperature [53,54], chemi-al dried with anhydrous sulphate [56] or lyophilised [57] beforeFE. Ling et al. [53] reported the extraction of several OCPsrom Chinese herbal medicines using SFE with CO2 at 25 MPand 50 ◦C (5 min static extraction time and 20 min dynamicxtraction time) using Florisil as trapping sorbent. A similarrocedure was used by Zuin et al. [54] for the determinationf OCPs and organophosphorus pesticides in Brazil’s medici-al plants. Mild extraction conditions (pure CO2; 10 MPa and0 ◦C, 5 min static plus 10 min dynamic extraction time) and18 as trapping adsorbent allowed for direct analysis of the
xtract by GC–ECD/Flame photometric detector (FPD) with norior cleaning procedure. Quan et al. [55] reported the extrac-ion of OCPs from ginseng by SFE using CO2 with 10 wt.%thanol–H2O solutions as modifier at 30 MPa, 60 ◦C and C18 as

tica C

teeuyeyit

eea[uPNeiveImaapscafaebsn

2

otanefittqots(oosas

sctitapa

pcptab

SbmMtabseaie

esea[MnaobucOiSvdo[ewpWt


rapping adsorbent. Pure CO2 was used by Zhu and Lee [56] toxtract PCBs from pine needles and exhibited good extractionfficiencies and recoveries (≥90%). The extract was collectedsing 3 mL of hexane and used directly in the GC–MS anal-sis. Crespo and Yusty [57] compared SFE with Soxhlet forxtracting PCBs from algae samples. Athough both methodsield comparable results, SFE offered the advantage of detect-ng all PCBs studied at lower concentrations, while extractionime and amount of solvents needed were reduced.

Several applications of SFE for extracting POPs from differ-nt animal tissues have also been reported [58–61]. Efficient SFExtraction of PBDEs from whale tissues was obtained with CO2t 40 ◦C and 28 MPa, flow rate 2 mL min−1 and trapping into C1858]. The extract was analysed directly without additional clean-p by GC–MS. The same SFE procedure was applied to extractCBs, chlordane, toxaphenes and PBDEs from seal tissue [59].erın et al. [60] made a comparison between Soxhlet and SFE

xtraction for the determination of OCPs and some metabolitesn frog tissues The study highlighted the main advantages of SFEersus Soxhlet procedures, including efficiency (higher recov-ries), time consumption, cost and more environment friendly.t must be kept in mind, however, that SFE requires an opti-ization in depth since the extraction behaviour is strongly

ffected by the type of sample. Similar to plant materials, inll the procedures reported above [58–60] animal tissue sam-les were desiccated with sodium sulphate before the extractiontep to make the sample matrix more accessible to the supercriti-al fluid. In fact, Antunes et al. [61] applied SFE to extract PCBsnd OCPs from fish muscle using three types of raw materials:resh fillet, fresh fillet grounded with anhydrous sodium sulphatend freeze-dried fillet. They found that supercritical CO2 couldxtract organochlorine compounds from freeze-dried fish fillets,ut they were not extracted efficiently from fresh fish. The pres-ure had a significant effect on extraction, while temperature didot significantly affect the efficiency of the extraction.

.4. Microwave-assisted extraction (MAE)

MAE uses microwave radiation (0.3–300 GHz) as the sourcef heating a solid sample–solvent mixture [62]. Due to the par-icular effects of microwaves on matter (namely dipole rotationnd ionic conductance) heating with microwaves is instanta-eous and occurs in the heart of the sample, leading to very fastxtraction. Heat generation in the sample by the microwaveseld requires the presence of a dielectric compound. The greater

he dielectric constant, the more thermal energy is released andhe more rapid is the heating for a given frequency. Conse-uently, the effect of microwave energy is strongly dependentn the nature of both the solvent and the solid matrix. Usually,he extraction solvent has a high dielectric constant, so that ittrongly absorbs the microwave energy. However, in some casesfor thermolabile compounds), the microwaves may be absorbednly by the matrix, resulting in heating of the sample and release
f the solutes into the cold solvent. Therefore, the nature of theolvent is of great importance in MAE: it should selectivelynd efficiently solubilize the analytes in the sample but, at theame time, it should absorb the microwaves without leading to a
Gbmc

himica Acta 590 (2007) 1–16 5

trong heating (so as to avoid eventual degradation of the analyteompounds). Thus, it is common practice to use a binary mix-ure (e.g. hexane–acetone, 1:1) where only one of the solvents absorbing microwaves. Other important parameters affectinghe extraction process are the applied power, the temperaturend the extraction time. Moreover, the water content of the sam-le needs to be carefully controlled to avoid excessive heating,llowing reproducible results.

The application of microwave energy to the samples may beerformed either in closed vessels with pressure and temperatureontrol (pressurised MAE) or in open vessels at atmosphericressure (focused MAE) [14,15]. Whereas in focused MAE,he temperature is limited by the boiling point of the solvent attmospheric pressure, in pressurised MAE the temperature maye elevated by simply applying adequate pressures [14].

Today MAE is considered a good alternative to traditionaloxhlet extraction for POPs in environmental solid samplesecause it reduces extraction time (e.g. 20–30 min per batch of asany as 12 samples), uses small amounts of solvents (30 mL inAE versus 300 mL in Soxhlet extraction) and improves extrac-

ion yields. Consequently, several applications are reported [63]nd an official EPA method 3546 (Microwave Extraction) haseen approved for the extraction of organic compounds fromolid environmental samples [2]. However, MAE has also sev-ral drawbacks. They include that the extract must be filteredfter extraction, polar solvents are needed, clean-up of extractss almost always needed (because MAE is very efficient) and thequipment is moderately expensive.

Several recent studies have reported the use of MAE forxtracting different POPs from plants [64–67] and animal tis-ues [68–72]. Since the water content of the sample has a greatffect on the extraction process, usually samples were air driedt room temperature [65–67], lyophilised [65,66], freeze-dried68] or chemical dried with anhydrous sulphate [70,71] before

AE. Cai et al. [64] used MAE to extract OCPs from Chi-ese teas before solid-phase microextraction (SPME)–GC–ECDnalysis. The recoveries of MAE were compared with thosef ultrasonic extraction and results showed that MAE providedetter recoveries (efficiencies) and shorter extraction times thanltrasonic extraction. Barriada-Pereira et al. [65] carried out aomparative study between MAE and Soxhlet extraction of 21CPs from plants using n-hexane–acetone (1:1, v/v) as solvent

n both cases. Both techniques showed similar recoveries butoxhlet extraction was more laborious and required higher sol-ent consumption and longer extraction times than MAE. Theeveloped MAE procedure was applied to the determinationf the same OCPs in tree leaves [66] and five species of plants67]. Carro et al. [68] reported a reliable and simple procedure toxtract PCBs from freeze-dried mussels using pressurised MAEith n-pentane–sodium hydroxide (5%) or dichloromethane–n-entane (1:1) at different temperature (70–90 ◦C) during 10 min.ittmann et al. [69] compared a combination of saponifica-

ion and liquid–liquid extraction (S-LLE) with MAE for the
C–ECD determination of trichlorobenzenes (TCBs) in cod. Inoth cases, n-pentane was used as the extraction solvent. Bothethods were appropriate for the detection of TCBs at con-
entration levels typically observed in marine biota. However,

6 tica Chimica Acta 590 (2007) 1–16

SsmaoMP1Hwlhoofcfcatvoo

2

(tiraitoiaiprmc

(liietc

bTtptp

F[

atPtuilssles

3mtytmrisvsf

rpfsPa(vci([


-LLE was more time consuming and required larger organicolvents volumes. Bayen et al. [70] reported the first validatedethod for the quantification of major PBDE congeners (47, 99

nd 100) in marine biological tissues using MAE with 25 mLf n-pentane–dichlorometane (1:1, v/v) as extraction solvent.AE was compared here to Soxhlet extraction for extracting

BDEs from standard reference materials (SRM 2978 and SRM588a) with comparable analytical results (<15% variation).owever, using MAE the extraction solvent volume (25 mL)as substantially lower than that required for conventional Soxh-

et extraction and the extraction time was reduced from severalours to 25 min. The method was applied to the determinationf POPs including PBDEs in a wide range of mangrove biotarganisms [71]. Pena et al. [72] developed a MAE procedureor the extraction of six regulated PAHs from polluted fish mus-le using n-hexane. Two types of fish samples representativesrom low and high fat content fish (turbot and salmon) wereonsidered for experimental optimization. Other samples of lownd high fat content (mussel and lamprey) were also analysedo verify the applicability of the developed procedure. Accuracyalidation using NIST SRM 2977 reference material was carriedut and recoveries around 90% for the studied compounds werebtained.

.5. Pressurised liquid extraction (PLE)

This technique, also named pressurized fluid extractionPFE), was originally launched by Dionex Inc. in 1995 underhe name accelerated solvent extraction (ASETM) [73,74]. PLEs a solid-liquid extraction process performed in closed-vessels atelatively elevated temperatures, usually between 80 and 200 ◦C,nd elevated pressures, between 10 and 20 MPa. Therefore, PLEs quite similar to SFE but CO2 is replaced by organic solventso mitigate potential polarity troubles [74]. Extraction is carriedut under pressure to maintain the conventional organic solventsn its liquid state, but extracting at temperatures well above theirtmospheric boiling points. Therefore, the solvent is still belowts critical conditions during PLE but has enhanced solvationower and lower viscosities and hence allows higher diffusionates for analytes. In this way the extraction efficiency increases,inimizing solvent needed and expediting the extraction pro-

ess.Both static and flow-through extraction systems can be used

Fig. 2) [14,75]: in the static extraction mode, the sample isoaded in an inert cell and pressurized with solvent heated abovets boiling point during some time (then, the extract is automat-cally removed and transferred to a vial). The “flow-throughxtraction mode” uses fresh solvent continuously introducedo the sample. This improves the extraction efficiency but, ofourse, diluting the extract [75].

In PLE, the pressure is of comparatively minor importanceecause its role is just to maintain the solvent in its liquid state.his reduces the number of parameters that need to be optimized

o achieve efficient extractions compared with SFE. The mainarameters to consider now are temperature and time and so theime devoted to development and optimization of the extractionrocedure can be reduced. Moreover, method set-up is gener-

fe1o

ig. 2. Schematic diagram of a PLE system. Reprinted with permission from10].

lly straightforward because the same solvent recommended inhe official and routine Soxhlet methods can be used. Therefore,LE is an attractive technique because it is fast (e.g. extraction

ime is approximately 15 min per sample), uses less solvent vol-me (15–40 mL), no filtration is required after extraction, thenstrumentation allows extraction in unattended operation (ateast 24 samples can be processed sequentially) and differentample sizes can be accommodated (e.g. 11, 22 and 33 mL ves-els are available). The main two disadvantages of PLE includeimited selectivity (so, it usually requires further clean-up of thextract obtained) and higher capital cost than SFE and MAEystems.

The acceptance of PLE as an official EPA method (method545) for the determination of POPs in a variety of environ-ental solid samples [2] has convinced many authors to use

his technique for sample preparation in environmental anal-sis [5,12,76]. Therefore, numerous applications of PLE forhe extraction of POPs from biota samples (vegetal and ani-

al) have been reported in the last 6 years [77–93]. In all theeported procedures PLE was applied to dry samples since, asn the case of Soxhlet extraction, the absence of water in theamples makes the sample matrix more accessible to organic sol-ents. Therefore, samples were dried by grinding with sodiumulphate [78,88] or with hydromatrix [80,82,90], air-dried [79],reeze-dried [83,85,86,89,90,93] or lyophilised [87] before PLE.

Weichbrodt et al. [77] used PLE for extracting organochlo-ine compounds from cod liver and fish fillets. Extraction waserformed with ethyl acetate–cyclohexane (1:1, v/v), allowingor direct use of gel-permeation chromatography as clean-uptep without solvent exchange. Suchan et al. [78] comparedLE with conventional Soxhlet extraction for extracting OCPsnd PCBs from fish fillets using two different solvent mixtureshexane–dichloromethane (1:1, v/v) and hexane–acetone (4:1,/v). It was found that PLE, with both solvents tested, wasomparable to Soxhlet. Tao et al. [79] applied PLE for extract-ng DDT and its metabolites from wheat with hexane/acetone1:1, v/v) (pressure 101 MPa, temperature 120 ◦C). Moreno et al.80] investigated the extraction of 65 pesticides including OCPs
rom greasy vegetable matrices such as avocado using PLE withthyl acetate–cyclohexane (1/1, v/v) at 120 ◦C and a pressure of2 MPa. Adou et al. [81] reported an analytical procedure basedn PLE before GC–ECD or GC–FPD for the determination of

tica C

daatsfawsual[tsiPrtmospgcmtaeoasstnntbsb1wasaPcd[

aPDsJOs

tta(etilolioph

deAataO

2

sowsec

puaiif[iatiobc

nast


ifferent pesticides in fruits and vegetables. The recoveries wereround 70% for almost all the compounds assayed. PLE waslso used by Haib et al. [82] for the extraction of OCPs fromobacco samples with acetone at 100 ◦C and 10 MPa of pres-ure. Focant et al. [83] used PLE to extract PCBs and PCDD/Fsrom highly fatty biological matrices (poultry, eggs, mackerelnd sperm whale blubber).The optimal extraction conditionsere obtained using hexane as extration solvent and a pres-

ure of 10 MPa. Similarly, Bjorklund et al. [84] reported these of PLE to extract PCBs from fat-containing matrices suchs fish meal and a certified reference material (CRM 349, codiver oil) using hexane as extraction solvent. Gomez Ariza et al.85] used a mixture of dichloromethane–pentane (15/85, v/v)o extract PCBs from biota samples (clams, oysters, eggs ofpoonbill, mussels and fish) using PLE. Quantitative recover-es (from 90 to 106%) were reported for both native and spikedCB congeners. The method was validated using the standardeference material SRM 2974 (mussel tissue) and comparedo Soxhlet and matrix solid-phase dispersion extraction. Kita-

ura et al. [86] developed a method based on the combinationf PLE with DMSO–acetonitrile (1:9, v/v) as the extractionolvent (14 MPa and 180 ◦C) and DMSO–acetonitrile–hexaneartitioning for the determination of PCBs and Dioxin con-ener (PCDDs/Fs) in lipid-rich biological matrices (beef, pork,hicken, and fish). Dioxin congener levels extracted by thisethod were almost identical to those obtained by conven-

ional solvent extraction methods, such as those employingcetone/hexane or toluene, but PLE was advantageous in short-ning analysis time. Martinez et al. [87] reported the extractionf PAHs from mussels by PLE with recoveries between 65nd 150% using hexane–dichloromethane (1:1, v/v) at a pres-ure of 10 MPa and a temperature of 150 ◦C. Janska et al. [88]elected hexane–acetone (1:1, v/v) as the most suitable extrac-ion solvent for extracting PAHs from fish tissue and spruceeedles samples using PLE. Relatively good recoveries (not sig-ificantly different from those obtained by employing classicechniques such as Soxhlet extraction and extraction enhancedy sonication) were obtained. Liguori et al. [89] also extractedeveral PAHs from blue mussel, salmon fillet, fish oil and fishy means of PLE using dichloromethane as solvent (pressure0 MPa, temperature 100 ◦C). Recoveries between 99 and 105%ere obtained for spiked samples. The method was validated by

nalysing two certified reference materials (CRM 2977, mus-el tissue and T0621, olive oil). PLE with dichloromethanes solvent (14 MPa and 100 ◦C) was also used for extractingBDEs from blue mussel [90] and PBDEs, hexabromo-yclododecane (HBCD) and methoxylated polybrominatediphenyl ether congeners (MeO-PBDEs) from sea lion blubber91].

In a recent investigation by Saito et al. [92] PLE waspplied with success to extract a broader spectrum ofOPs (PCBs, PBDEs, OCPs, etc.) from biological tissues.ichloromethane–acetone (1:1, v/v) was used as the extraction

olvent at a temperature of 100 ◦C and a pressure of 10 MPa.iang et al. [93] reported a PLE procedure to extract also severalPPs (PCBs, PCDDs/DFs and PCNs) from freeze-dried seafood

amples (fish, bivalves, shrimp, crab, and cephalopods).

ofib

himica Acta 590 (2007) 1–16 7

When water is employed as the extraction solvent in PLEhe authors tend to use a different name to highlight the facthat water is an environment-friendly solvent. Thus, terms suchs pressurized hot water extraction, subcritical water extractionSWE), superheated water extraction and high temperature waterxtraction can be found in the literature [12,76,94]. Becausehe polarity of water decreases markedly as the temperature isncreased, superheated water at 100–200 ◦C, under a relativelyow pressure, can act as a medium to non-polar solvent (ethanolr acetone) and is an efficient extraction solvent for many ana-ytes [12,76,94]. A limitation in extracting with hot water is thatt fails to recover compounds that are hydrophobic, thermolabile,r easily hydrolysable. Therefore, until now, only a few exam-les of subcritical water extraction of POPs from biota samplesave been published [76].

Recently, Morales et al. [95] used water modified with sodiumodecyl sulfate (SDS) as extractant for extracting PAHs fromnvironmental solid samples (soil, sediment, trout and sardine).comparison between three operational modes (static, dynamic

nd static–dynamic mode) was carried out and efficiencies closeo 100% were obtained for the three modes tested. Wennrich etl. [96] also applied subcritical water extraction (SWE) to extractCPs and chlorobenzenes from fruit and vegetables.

.6. Matrix solid-phase dispersion (MSPD)

MSPD is a process for the disruption and extraction ofolid samples introduced in 1989 [97]. MSPD combines aspectsf several analytical techniques, performing sample disruptionhile dispersing the components of the sample on and into a

olid support. In this way a chromatographic material is gen-rated that possesses unique character for the extraction ofompounds from the dispersed sample [98].

In MSPD the sample is mixed (for liquid and semi-solid sam-les) or blended (for solid samples) with an appropiate sorbentntil a homogeneous mixture is obtained (complete disruptionnd dispersion of the sample on the solid support); this mixtures packed into an empty column, from which the analytes ofnterest are eluted with a suitable organic solvent while inter-ering matrix compounds are selectively retained on the column99]. Another possibility is the elution of interfering compoundsn a washing step while, the target analytes are next eluted bydifferent solvent. Finally, additional clean-up is performed or

he sample is directly analysed. Sometimes, the MSPD columns coupled on-line with a solid phase extraction (SPE) columnr, as in several applications, the SPE sorbent is packed in theottom part of the MSPD column to remove interfering matrixomponents [97,100].

MSPD can be regarded as a valid sample preparation tech-ique, alternative to more classical methods, especially for solidnd semisolid samples. It is simple, requires a small sampleize, has a short extraction time, uses less solvent than conven-ional techniques, does not require preparation and maintenance
f equipment and offers the possibility of simultaneously per-orming extraction and cleanup. However, the negative aspects that MSPD is fairly labour intensive, requiring the sample toe ground up with the solid matrix and packed into a column

8 tica C

fv

buisabatepbit

iHPfiaPehMsw

fbatUCcwAepwdcSmfs[fUao1eots

owafanagsilac

3

cfmlgcbcordooo

3

ieapcbdusc(pt3Co


or extraction, and quite a number of applications still use largeolumes of solvents for extraction and clean-up.

The selectivity of a MSPD procedure depends on the sor-ent/solvent combination used. Most methods reported to datese reverse-phase materials, such as C8- and C18-bonded sil-ca as the solid support; silica, Florisil and chemically-modifiedorbents are used less frequently. For analyte extraction fromnimal tissues, C18-bonded silica is by far the most popular sor-ent while, for plant samples, both C8- and C18-bonded silicand also Florisil are used extensively [98]. The nature of the elu-ion solvent is also important since the target analytes should befficiently desorbed while the bulk of the remaining matrix com-onents should be retained in the column. Most sorbents haveeen tested in combination with a large variety of solvents, rang-ng from alkanes through toluene, dichloromethane and alcoholso water at elevated temperatures.

Most environmental applications of MSPD deal with extract-ng pesticides from fruit, vegetables and animal tissue [98].owever, there are also papers reporting the extraction of otherOPs such as PAHs from fish tissue [101], PCBs from shell-sh and fish [85], PCBs from animal fatty samples [102], PCBsnd PBDEs from different biota material [103]; and pesticides,CBs, PBDEs and polybrominated biphenyl (PBBs) from sev-ral marine species [104]. Dry samples are most effectivelyomogenised and dispersed with the solid support used inSPD. Therefore, samples are usually dried with anhydrous

odium sulphate [101–103] or freeze-dried [85] before blendingith the MSPD sorbent.Pensado et al. [101] described the performance of MSPD

or the extraction of PAHs from fish tissue (salmon and tur-ot). The suitability of different solid supports (Florisil, C18,nd acidic silica gel) was tested as well as the influence onhe extraction efficiency of the natural fat content in samples.nder optimal conditions, tissue samples were dispersed with18 and anhydrous sodium sulphate and transferred to a SPEartridge containing florisil and C18. Cartridges were elutedith acetonitrile and the extract directly analysed. Gomez-riza et al. [85] compared MSPD to PLE and Soxhlet for

xtracting PCBs from shellfish and fish. In MSPD, the sam-les (freeze-dried) were blended with Florisil, and the mixtureas placed in a glass column containing Florisil and eluted withichloromethane–pentane (15:85, v/v). The obtained extract waslean enough for direct analysis by GC–MS and GC–ECD.imilar advantages were obtained using both MSPD and PLEethods, but the capital cost of MSPD is much lower than that

or PLE and Soxhlet. Different combinations of normal phaseorbents and elution solvents were evaluated by Criado et al.102], in terms of extraction yield and lipids removal efficiency,or the isolation of PCBs from butter, chicken and beef fat.nder optimal conditions, the sample was dispersed on Florisil

nd transferred to the top of a solid-phase extraction cartridgef Florisil. Non-coplanar PCBs were quantitatively eluted with5 mL of n-hexane. Coplanar and non-coplanar congeners were
luted with n-hexane–dichloromethane (90:10). In this case, 1 gf acidic silica was also placed in the bottom of the SPE car-ridge, in addition to the 5 g of Florisil. With this combination oforbents, the small amount of fat which elutes from Florisil was
cd(o

himica Acta 590 (2007) 1–16

xidized by the silica layer, leading to cleaner extracts, whichere directly analysed by GC–MS and GC–ECD. Martınez et

l. [103] reported the extraction of PCBs and PBDEs from dif-erent biota material (beef fat, chicken fat, turbot fish musclend dogfish liver) using a MSPD extraction cartridge of C18 and-hexane as elution solvent. PCBs and PBDEs were fraction-ted on a second cartridge containing silica. The same researchroup [104] developed a multiresidue method for the trace analy-is of 15 OPPs from several pesticides, PBDEs, PCBs and PBBsn marine species (turbot, clam, mussel and cockcle). The ana-ytical procedure is based on MSPD of the sample using C18s sorbent and basic alumina, in the bottom part of the MSPDolumn to clean-up with subsequent elution with hexane.

. Extract clean-up

Whichever technique is used for extraction, various matrixomponents such as lipids, carotenoids, pigments and resins arerequently present in the extract and must be eliminated to per-it a more definitive identification and quantification of lower

evels of analyte and to minimize deterioration of chromato-raphic performance. Thus, the removal of co-extracted matrixomponents is critical and so different clean-up procedures haveeen developed to minimise their negative effects. Moreover, thelean-up step is usually necessary to remove not only the bulkf the co-extracted material, but also those compounds closelyelated to the analytes that could potentially interfere in the finaletermination. In this latter case adequate separation schemesr fractionation processes to allow for isolation of sub-groupsf compounds (fractionation of the extract into different classesf compound) has to be carried out.

Let us revise the most common strategies for extract clean-up.

.1. Classical liquid adsorption chromatography

Classical liquid adsorption chromatography is still the dom-nant technique for purification and fractionation of biotaxtracts. This classic technique is used in an “off-line” modend involves passing extracts through several adsorbent columnsrepared in the laboratory or through solid-phase extractionartridges. Liquid adsorption chromatography can discriminateetween the target compounds and the matrix components to aegree that depends on the selectivity of the sorbent (sorbents)sed. Alumina, silica gel and Florisil columns in different meshizes, levels of activity and column sizes (either separately or inombination) are widely used. Sometimes, an alkaline treatmentsaponification) or a treatment with sulphuric acid is necessaryrior to, or in conjunction with, adsorption columns to removehe bulk of co-extracted lipids (Table 1). Official EPA Methods630C, 3610B and 3620B (using silica gel, alumina and Florisilleanup, respectively) have been approved for the purificationf organic extracts from solid environmental samples [2].

Pena et al. [72] reported the use of silica gel cartridges to
lean-up fish tissue extracts containing PAHs after alkaline lipidsigestion. Similarly, extracts of different vegetables and fruitslettuce, tomato, cabbage, apple, grape and pear) were purifiedn a deactivated silica gel (15% water) column before PAHs

N. Fidalgo-Used et al. / Analytica Chimica Acta 590 (2007) 1–16 9

Table 1Methods for the clean-up of POPs in biota extracts: classical liquid adsorption chromatography

Compound Matrix Adsorption column Ref.

PAHs Fish tissues Silica cartridges after alkaline digestion [72]PAHs Vegetables and fruits Silica [105]PCBs Seaweeds Silica [57]PCBs, OCPs, PAHs Mussels Silica [106,107]OCPs Frogs Silica [60]PBDEs Mussel, eel, porpoise and

cormorant tissuesSilica after sulphuric acid treatment [32]

PCBs, OCPs, OPPs Whale tissues Automatic LC silica [108]PCBs, OCPs Fish tissues Silica impregnated with sulphuric acid [31]PCBs, DDTs Longfin eel tissues Acidic Silica + basic silica [41]PAHs Fish liver Alumina [109]PAHs Mussels Alumina after alkaline digestion [87]PAHs Lichens Alumina [52]PAHs, OCPs, PCBs Fish liver Alumina [22]PBDEs Fish muscle, shellfish and fish

liverAlumina [34]

OCPs Vegetables and sunflowers seeds Florisil [110,111]DDTs Plants Florisil after sulphuric acid treatment [79]PCBs Clam, spoonbill eggs, fish,

mussel and oysterFlorisil after sulphuric acid treatment [85,112]

PCBs, OCPs Fish liver, blubber whale andblack scabbardfish

Florisil [61,113,114]

PAHs Plants, mussel, eel and fish tissues Florisil [17,48,115,116]OCPs Tree leaves Carbon cartridges [66]PAHs Pine needles Alumina cartridges [49]PAHs, PCBs, DDTs Mussel and krill Silica + alumina (3:1) [117]POPs Cod liver Florisil + carbon + basic alumina [118]tricholorobenzenes Fish tissues Alumina + acidic silica [69]PCBs Mussels Silica + alumina [119]PCBs Fish tissues Sulphuric acid + silica + silver nitrate coated

silica + Florisil[37]

PCBs Fish tissues Neutral silica + acidic silica + ENVI-Carb cartridge [24]OCPs Tobacco Florisil + silica [82]OCPs Vegetation samples Alumina + Florisil cartridges [65,67]OCPs Fish tissues Alumina + silica + florisil impregnated with KOH [31]OCPs, PCBs Silverfish and krill Neutral silica + acidic silica [23]PBDEs Mussel, eel, porpoise and

cormorantAlumina followed by silica [32]

PBDEs Cormorant liver Alumina followed by silica after sulphuric acidtreatment

[35]

PBDEs Whale tissues Acidic silica + neutral silica + basic silica [120]PBDEs Whitefish and rainbow trout Acidic silica + neutral silica + basic silica followed

by alumina[121]

PBDEs, PCDDs/Fs, PCBs, PBBs Horse mackerel tissues Acidic silica + neutral silica + basic silica [122]PCBs, PCDD/Fs, PCNs Sea food Acidic silica + neutral silica + basic silica [93]PCBs, PCDD/Fs, PCNs Pine needles Acidic silica + neutral silica + basic silica [44]OCPs, PCBs, PBDEs Guillemots liver and guillemots

eggsAlumina + silica + acidic silica [123]

PBDEs, Fish and blue mussels tissues Alumina + silica + acidic silica [33]PCBs, PBDEs Ringed seal blubber Alumina + silica + acidic silica [124]PCBs, PCDDs/Fs Lipid biota matrices Silica + carbon [86]PCBs, OCPs, PCDDs/Fs Bald eagles tissues Silica + alumina + acidic silica + carbon [125]PCBs, PCDDs/Fs, PBDEs Fish tissues Silica + alumina + acidic silica + carbon [46]PBDEs, PCBs, PCDDs/Fs Beef fat Automated acidic silica + neutral silica + basic [83,126]

dgoie

f
etermination [105]. Crespo and Yusty [57] used also a silica el column to clean-up seaweed extracts in the determinationf PCBs. A column filled with activated silica gel was used forsolation of fractions containing PCBs and OCPs from musselxtracts [106] and fractions containing PCBs, OCPs and PAHs
[vst

silica followed by basic alumina and carbon

rom extracts of the same type of samples [107]. Nerın et al.
60] tested several adsorbents (3% deactivated silica, 5% deacti-ated Florisil) as well as their combinations and different elutionolvents (in volume and nature) to clean-up frog extracts forhe determination of OCPs. Best results were obtained with 3%

1 tica C

dbifumsgcOsrd

lppdot[e

sAFe[oatams

cdltmaenotcsmtc(o

(seb

opdwcns(tdo[omi[

iwclbstuapwA/tbwsPe[awvwe[Cbmmearatfi

0 N. Fidalgo-Used et al. / Analy

eactivated silica and n-hexane as eluent. Extracts of differentiota samples (mussel, eel, porpoise and cormorant) contain-ng PBDEs were also clean up on a silica gel column afterat destruction with sulphuric acid [32]. Also, automatic liq-id chromatography on a silica gel column, using n-hexane asobile phase, was reported to clean-up extracts of whale tis-

ues in the determination of several POPs [108]. Activated silicael impregnated with concentrated sulphuric acid was used forlean-up extracts of fish species containing PCBs and acid-stableCPs [31], while a column with two layers of silica, treated with

ulphuric acid and potassium carbonate, respectively, was alsoeported to be useful to clean-up longfin eels extracts for theetermination of PCBs and DDTs [41].

Vives et al. [109] used an alumina column to clean-up fishiver extracts. Elution with n-hexane–dichloromethane (1:2, v/v)rovided the PAHs fraction. Martınez et al. [87] used alumina forurification of mussels extracts containing PAHs after alkalineigestion. Alumina columns were also applied for purificationf lichens extracts containing PAHs [52] for isolation of frac-ions containing PAHs, OCPs and PCBs from fish liver extracts22] and for clean-up fish muscle, shellfish tissue and fish liverxtracts when determining PBDEs [34].

For the clean-up of OCPs in fresh vegetables [110] andunflower seeds [111] extracts a Florisil column was reported.lso, plant extracts containing DDTs were purified by using alorisil column after sulphuric treatment [79]. Florisil was alsomployed to clean-up mussels extracts after sulphuric treatment112] and different biota (clam, spoonbill eggs, fish, mussel andyster) extracts [85], prior to the determination of PCBs. Usinggain a Florisil column the separation of PCBs and OCPs frac-ions on extracts of fish liver samples [113], whale blubber [114]nd black scabbard fish [61] and clean-up of PAHs in plant [115],ussel [48], eel, goldfish, catfish [17] and in other several fish

pecies extracts [116have been reported.On the other hand, Barriada-Pereira et al. [66] compared

artridges filled with four different sorbents: Florisil, a tan-em of Florisil and alumina, silica, and carbon to clean-up treeeaves extracts prior to OCPs determinations. Carbon provedo be the sorbent, providing colourless eluates, cleaner chro-

atograms and lesser interferences. Similarly Florisil, silicand alumina cartridges as well as glass columns (filled withither Florisil, silica or alumina) were also compared for pineeedles extracts purification previous to final determinationf PAHs and alumina disposable cartridges were chosen ashe most efficient [49]. Silica gel, alumina, aminopropyl-silica,yanopropyl-silica, Florisil, graphitized nonporous carbon andilica gel–alumina mixture (3:1) were used for column chro-atographic clean-up of PAHs, PCBs and DDTs in mussel

issue and krill samples in combination with modified super-ritical CO2 as the mobile phase [117]. A silica gel–alumina3:1) column was shown to offer the best performance in termsf clean-up efficiency [117].

Sometimes it may be necessary to use more than one column
adsorbent) to obtain adequate clean-up and/or fractionation ofample extracts. In this vein, extracts of cod liver containing sev-ral POPs were fractionated with Florisil, activated carbon andasic alumine column chromatography [118]. A combination
bMt

himica Acta 590 (2007) 1–16

f 5% deactivated alumine and silica impregnated with sul-huric acid (acidic silica) was used as a clean-up column in theetermination of trichlorobenzenes in fish tissues samples [69],hile mussel extracts were cleaned-up using a silica–alumina

olumn prior to PCBs determination [119]. Similarly, a combi-ation of sulphuric acid treatment, silica gel, silver nitrate coatedilica gel and Florisil was used for purification extracts of fishmullet) containing PCBs [37]. A multilayer silica column (neu-ral/acidic) was recommended to clean-up fish extracts for PCBseterminations. That clean-up was followed by a separation ofrtho-PCBs from non-ortho-PCBs on a ENVI-Carb cartridge)24]. Florisil followed by silica gel clean-up was used for extractsf tobacco) containing low polar OCPs [82]. 5% deactivated alu-ina was added to a Florisil cartridge for purification of OCPs

n different vegetation samples (grass, five species of plants)65,67].

A column made with superposed layers of alumina, sil-ca and Florisil impregnated with KOH and elution was doneith n-hexane–dichloromethane (3:1, v/v) [31] was used for the

lean-up of non-acid stable OCPs in fish extracts) while a multi-ayer silica column (neutral/acidic) was applied for fractionationetween PCBs and OCPs in extracts of krill and silverfishamples [23]. Two separated columns, one packed with 5% deac-ivated alumine and the other with 3% deactivated silica weresed sequentially to clean biota samples (mussel, eel, porpoisend cormorant) extracts containing PBDEs [32] and a similarrocedure, but after an additional treatment with sulphuric acid,as used for PBDEs clean-up in cormorants liver extracts [35].multilayer silica column (acidic silica /neutral activated silica

basic silica) was proposed for the clean-up of PBDEs in whaleissue extracts [120], while this silica column treatment followedy a basic alumina column was used for extracts purification ofhitefish and rainbow trout containing PBDEs [121]. Multilayer

ilica columns were also reported to clean-up PBDEs, PCDD/Fs,CBs and polybrominated biphenyls (PBBs) in horse mackerelxtracts [122] and PCBs, PCDD/PCDFs and PCNs in seafood93], and pine needle extracts [44]. The clean-up of OCPs, PCBsnd PBDEs in extracts of guillemots livers and guillemot eggsas carried out on a multilayered column packed with deacti-ated alumina, activated silica and activated silica impregnatedith sulphuric acid [123]. The same multilayer column was

mployed to clean-up PBDEs in fish and blue mussels extracts33], and PBDEs and PCBs in ringed seal blubber extracts [124].oplanar-PCBs, PCDDs and PCDFs in extracts of lipid-richiological matrices were fractionated by resorting to tandemultilayer silica gel-activated carbon (MLS-AC) column chro-atography [86], while PCBs, OCPs and PCDD/DFs in bald

agle tissue extracts were sequentially subjected to silica gel,lumina, acidic silica and activated carbon column chromatog-aphy for adequate clean-up and fractionation [125]. Isosaari etl. [46] used a similar procedure for adequate clean-up and frac-ionation of PCBs, PCNs, PCDD/DFs and PBDEs in extracts ofsh tissue.

A commercially available automated clean-up procedureased on multi-column clean-up system (Power-Prep, Fluidanagement Systems, Waltham, MA, USA) was evaluated for

he determination of PBDEs, PCBs, PCDDs and PCDFs in for-

N. Fidalgo-Used et al. / Analytica Chimica Acta 590 (2007) 1–16 11

n-up

tamcuwVstsc

as

3

r

TM

C

PPPPPPPOPPPPOOPPPPP

Fig. 3. Schematic diagram of the automated clea

ified beef fat as quality control samples [83]. The commerciallyvailable clean-up system (Fig. 3) based on the use of disposableultilayer silica (acidic, basic and neutral), basic alumine and

arbon columns can clean-up ten extracted samples containingp to 1 g of lipids in less than 2 h. The system is made of three-ay and six-way electrostatic valves driven by PC software.alve modules (V1–V6) are responsible for the selection of the
olvent and columns. The fractionation procedure can be tunedo allow the operator to collect different fractions at differentteps of the purification process [83,126,127]. When the extractontains more than 1 g of fat, a high capacity silica column (28 g
smmg

able 2ethods for the clean-up of POPs in biota extracts: gel-permeation chromatography

ompound Matrix

CBs, OCPs Fish tissuesAHs Spruce needles and fish tissuesCBs Coots tissuesesticides Animal fatAHs VegetationCBs, PBDEs, PCNs MusselsCBs, OCPs Guillemot, mussels, green sea urchins, sculpin andCPs Burbot and troutCBs, OCPs SalmonBDEs Herring gull eggsolyciclic musks Marine mammals tissuesCBs, OCPs Perch and seal blubberCPs, PCBs, PBDEs BiotaCPs, PCBs, PCDD/Fs Fish and whale tissuesCBs, OCPs, MeSO2-PCBs Polar bear adipose tissueCBs, OCPs Fish tissuesBDEs Dolphin and porpoise tissuesOPs Biological tissuesBDEs Trout

system. Reprinted with permission from [126].

cidic, 16 g basic, 6 g neutral) is located before the multilayerilica column (in order to remove a large amount of fat).

.2. Gel-permeation chromatography (GPC)

This technique also referred to as size exclusion chromatog-aphy (SEC) and based on molecular size separation. GPC
eparation is used primarily to fractionate and remove lipidicaterial (>500 A), which elute first from the column in biotaatrices. Most workers used Bio-Beads S-X3 (200–400 mesh)
els in a range of column size and of mobile phases (Table 2).

GPC column Ref.

Biobeads S-X3 [78]Biobeads S-X3 [88]Biobeads S-X3 [38]Biobeads S-X3 [128]Biobeads S-X3 after Alumina + silica column [51]Silica + Biobeads S-X3 + alumina column [45]

clams Biobeads S-X3 after Florisil column [129]Biobeads S-X3 after Florisil column [130]Biobeads S-X3 after Florisil column [131]Biobeads S-X3 after Florisil column [132]Biobeads S-X3 after Florisil column [30]Biobeads S-X3 after silica column [133,134]Biobeads S-X3 after silica column [19]Biobeads S-X3 after silica column [18]Biobeads S-X3 after silica and Florisil columns [135]Biobeads S-X3 after alumina column [136]Biobeads S-X3 after silica column [42]Biobeads S-X3 after silica column [90]Biobeads S-X3 after alumina column [137]

1 tica C

Sasnemcoi

CeeeebuAwtae

ttaaBefcmsaAeegm[gbewwcpfOou[iaeuc

pCcAne[tM[c

3

ecseiafig

cueiii

vwts[

bmHpa[A

pSleSsSfi


everal key advantages of GPC, over other methods currentlyvailable for the clean-up of lipids include unlike typicalaponification or concentrated sulphuric acid treatments it ison-destructive, it allows handling larger masses of lipids inach sample (e.g. compared to adsorption columns) and GPC isore applicable than adsorption chromatography to “unknown”

ontaminants isolation when little information on the polarityr chemical functionality of the molecule is available. Finally,t can be fully automated.

In this vein, an official EPA method (Method 3640A GPCleanup) has been approved for the purification of organicxtracts from solid environmental samples [2], while Suchant al. [78] reported the purification of PCBs and OCPs in fishxtracts by GPC on a S-X3 Biobeads column with cychlohexane-thyl acetate (1:1, v/v) as mobile phase. The same GPC columnut with chloroform as mobile phase was used for the clean-p of PAHs in spruce needles and fish tissue extracts [88].lso, a S-X3 Biobeads column was used by Dodder et al. [38]ith cyclohexane–dichloromethane (60:40, v/v) as eluent for

he clean-up of PCBs in extracts of coots tissues, and with ethylcetate–cyclopentane (70:30, v/v) for clean-up of animal fatxtracts containing pesticides [128].

One main disadvantage of the GPC system is that it is difficulto completely remove all traces of the lipids. Therefore, fur-her clean-up steps are often necessary by resorting to the liquiddsorption chromatography columns mentioned above. Smith etl. [51] used a column of alumina and silica gel before GPC onio-Beads S-X3 for the clean-up of PAHs in pasture vegetationxtracts while Pan et al. [45] employed a multilayer silica columnollowed by a Bio-Beads S-X3 column and finally and aluminaolumn for clean-up of PCBs, PCNs and PBDEs in extracts ofussel. PCBs and OCPs in extracts of biota (guillemot, mus-

els, green sea urchins, sculpin and clams) were purified usingBio-Beads S-X3 column followed by a Florisil column [129].similar procedure was employed for the clean-up of OCPs in

xtracts of trout and burbot samples [130], OCPs and PCBs inxtracts of salmon samples [131], PBDEs in extracts of herringull eggs [132] and polycyclic musk fragrances in extracts ofarine mammals tissue (bear, seal, sea lion, sea otters, dolphins)

30]. A Bio-Beads S-X3 column followed by a column of silicael was used for clean-up PCBs and OCPs in perch [133] and seallubber extracts [134], OCPs, PCBs and PBDEs in several biotaxtracts [19] and OCPs, PCBs and PCDD/Fs in fish and bowheadhale tissue extracts [18]. This combined GPC-silica procedureas combined with a Florisil column for isolation of fractions

ontaining PCBs, OCPs, and MeSO2-PCBs in polar bear adi-ose tissue extracts [135], while GPC on Bio-Beads S-X3, butollowed by alumina column was also used for the clean-up ofCPs and PCBs in fish tissue extracts [136]. PBDEs in extractsf dolphin and porpoise blubber, liver and kidney were cleaned-p by GPC (Bio-Beads S-X3) followed by activated silica gel42] the same procedure and was used for purification of POPsn extracts of biological tissues [90] or followed by an activated
lumina column, treatment for purification of PBDEs in troutxtracts [137]. Nowadays, high efficient GPC columns are alsosed for previous clean-up of OPPs in extracts of biota in alassical HPLC system. Moreno et al. [80] evaluated the com-
mcmi

himica Acta 590 (2007) 1–16

arative performance of Envirogel GPC columns (Waters) with18, Florisil and alumina cartridges for purification of pesti-ides in avocado. It was found that GPC offers the best results.lso, two Envirosep-ABC columns (Phenomenex) were con-ected in series for purification of pesticides in fatty matricesxtracts, using acetate–cyclohexane (1:1, v/v) as mobile phase138]. This high performance GPC column was compared withhe classical Bio-Beads S-X3 column for the purification of

eSO2-PCBs and OCPs in extracts of marine mammal tissues139]. The best results were obtained when the classical GPColumn was employed.

.3. Solid-phase microextraction (SPME)

Solid-phase microextraction (SPME) is a relatively recentxtraction technique [140,141], mainly suited for aqueous matri-es, based on the partition of target analytes between a polymerictationary phase coating a fused silica fiber, and the samplextract. During extraction the coated fiber is either directlymmersed into the liquid extract or exposed to the headspacebove the liquid. After extraction, the analytes, retained in theber, are thermally desorbed in the injector of a gas chromato-raph for GC analysis.

SPME represents today a convenient alternative to moreonventional extraction methods of sample preparation of liq-id samples (i.e. liquid-liquid extraction (LLE) and solid-phasextraction (SPE). SPME eliminates the use of organic solvents,s significantly quicker and simpler than both LLE and SPE andntegrates extraction, preconcentration and sample introductionnto a single step [142–144].

Since its introduction in 1990 [141] to analyse relativelyolatile compounds in the environmental field, SPME has gainedidespread applications for determination of organic pollu-

ants, including POPs, in different types of samples (water,oil, food and biological fluids) as reported in several reviews142,145–147].

Water is by far the most widely analysed type of sampley SPME–GC. This is due to the fact that SPME application toore complex matrices, such as biota, is not so forward straight.owever, SPME can be used as a simple clean up/enrichmentrocedure for POPs determinations in vegetal and animal tissuesfter previous extraction by liquid–solid extraction, as Soxhlet39], ultrasonic assisted extraction [148,149], MAE [64,150],SE [96] or SFE [151].Fidalgo-Used et al. [39] developed a clean-up/enrichment

rocedure for OCPs in fish tissue samples based onoxhlet extraction of the OCPs from the sample fol-

owed by SPME–GC–ECD over the corresponding organicxtract. A representative chromatogram corresponding to thePME–GC–ECD analysis from a fish tissue organic extractpiked at 1 ng mL−1 level in the extract (final concentration in thePME vial of 0.1 ng mL−1) and its respective blank (unspikedsh tissue extract) it shown in Fig. 4. A combined analytical
ethod involving sequential ultrasonic extraction, sulfuric acid
lean-up and headspace SPME–GC–MS was used for the deter-ination of PCBs [148] and OCPs [149] in bird liver. SPME–GC

n combination with MAE for the determination of OCPs in

N. Fidalgo-Used et al. / Analytica C

Fig. 4. Chromatograms obtained by SPME–GC–ECD of: (a) fish tis-sue extract spiked with OCPs (final concentration in the SPME vial of100 ng L−1) and (b) unspiked fish tissue extract. Peak assignment: (1)hexachlorobenzene (HCB), (2) �-hexachlorocyclohexane (�-HCH), (3) �-HCH, (4) �-HCH, (5) �-HCH, (6) heptachlor, (7) aldrin, (8) isodrin, (9)dichlorodiphenyldichloroethylene (p,p′-DDE), (10) endosulfan �, (11) dieldrin,(fR

mmdss3t

fvA

bl(TSaI

4

itt

sswoiFbmeafmsStadaiseweof0nuntaPttby on-line clean-up with sorbents such as Florisil, silica gel,alumina, 2,3-dihydroxypropoxypropyl (Diol) and cyanopropyl(CN) in the extraction cell has been reported by Gomez-Arizaet al. [85]. The best results were obtained using Florisil to retain

12) endrin, (13) dichlorodiphenyldichloroethane (p,p′-DDD), (14) endosul-an �, (15) dichlorodiphenyltrichloroethane (p,p′-DDT), (16) methoxychlor.eprinted with permission from [39].

edicinal plants [150] and a novel fiber coating of polyphenyl-ethylsiloxane (PPMS) was also combined with MAE for the

etermination of OCPs in Chinese teas [64]. That novel porousol–gel PPMS fiber was reported to offer higher sensitivity andelectivity for OCPs compounds, higher thermal stability (up to50 ◦C) and longer life time (adequate use for more than 150imes) than commercial polydimethylsiloxane (PDMS) fibers.

On the other hand, Wennrich et al. [96] used SPME–GC–MSor the determination of OCPs and chlorobenzenes in fruit andegetables after a pre-extraction of analytes from the sample bySE.Finally, Rodil et al. [151] have developed a new approach,

ased on simultaneous SFE-sample clean-up using SPE fol-owed by SPME–GC–MS, for the determination of several POPsOCPs, PCBs, PBBs and PBDs) in cultured marine species.he influence of several parameters in the efficiency of thePE/SPME combination was investigated by chemometricspproaches and the proposed procedure was validated withAEA 406 reference material analysis.

. Integrated extraction and clean-up

Several researchers have examined the suitability of integrat-ng the clean-up step into SFE or PLE techniques by resortingo the use of sorbents in the extraction cell which would retainhe matrix components (trapping sorbents). In the case of biota

Fb

himica Acta 590 (2007) 1–16 13

uch sorbents are used mainly to retain the sample fat. However,o far relatively few publications have been published dealingith on-line combined extraction and clean-up procedures basedn SFE [53,59,152] or PLE [84,85,89,153,154] treatments. Fornstance, Ling et al. [53] investigated different sorbents (e.g.lorisil, C18, silica gel and neutral alumina) as “trapping” sor-ents in SFE for the determination of OCPs in chinese herbaledicine. Florisil was found to provide the most facile and

ffective integrated clean-up. Two fat retainers, basic aluminand Florisil, were assayed for the lipid-free extraction of PCBsrom a model fatty sample using SFE [152] where basic alu-ina was finally utilized to selectively clean-up PCBs in the fat

ample extract. Basic alumina was also used as fat retainer inFE for the determination of OCPs, PCBs and PBDEs in seal

issue [59]. Bjorklund et al. [84] evaluated the ratios between fatnd fat retainer to obtain fat-free extracts. They assayed for fiveifferent fat retainers, including Florisil, basic alumina, neutrallumina, acidic alumina and sulphuric acid-impregnated silican order to clean-up PCBs in fish using PLE and the use ofulphuric acid-impregnated silica provided the cleanest PCBsxtracts. Sulphuric acid-impregnated silica as fat retainer in PLEas also successfully used for the determination of PCBs in sev-

ral fat containing matrices including lard fat, pork fat, cod liveril, fish meal, feed poultry and vegetable feedstuff. Four dif-erent fat/fat retainer ratios (FFRs) were tested (0.100, 0.075,.050 and 0.025) at 50 and 100 ◦C using n-pentane, n-hexane or-heptane as extraction solvent. No fat was co-extracted whensing a FFR ratio of 0.025 and the chromatograms showed veryice base-lines as seen in Fig. 5 [153]. In another approach,he PLE cell was filled up with aluminium oxide, silica gelnd magnesium sulphate for the extraction and purification ofAHs in blue mussel and salmon [89], but a further purifica-ion by GPC was required. PLE extraction of PCBs from biotaissues (clams, oysters, eggs of spoonbill, mussels and fish)

ig. 5. Influence of the amount co-extracted fat on the chromatographicehaviour of PCBs. Reprinted with permission from [153].

1 tica C

cpiuP

5

ctspcatierutracHtcts

oaa

A

A(sd

R


oextracted lipids from the matrix. The resulting extract, afterre-concentration, turned out to be clean enough for its directnjection into GC–MS and GC–ECD [85]. Eljarrat et al. [154]sed alumina as fat retainer in PLE for the determination ofBDEs and HBCD in fish tissue.

. Conclusions

Sample preparation is still the most critical and time-onsuming step within the overall analytical procedure neededo obtain accurate determination of POPs in environmental biotaamples. Due to the peculiarities of these samples (especially theresence of fat and the presence of analytes at trace/ultratraceoncentrations) selection of the appropriate techniques to purifynd preconcentrate the analytes, along with a careful optimiza-ion of the corresponding operational parameters, are the mostmportant aspect to care of. As shown in this review, sev-ral alternative extraction techniques have been developed toeplace tedious conventional methodology (e.g. Soxhlet) oftensed in official analytical procedures. Among the new extractionechniques, SFE and PLE offer the advantages of significantlyeducing the amount of organic solvent consumed and easyutomation. Moreover, they also provide the possibility for effi-ient and fast on-line clean-up, by using selective trapping.owever, the comparatively high investment cost of these new

echniques explains that conventional Soxhlet extraction, inombination with adsorption columns and/or GPC for purifica-ion and fractionation of extracts, is still widely used being theample preparation reference method in numerous applications.

Finally, emerging techniques such as MSPD or SPME havenly been used in a limited number of applications so far. Inny case, the results obtained are promising and show that theirpplication should more likely expand in the near future.

cknowledgments

The autors are grateful to the “Fundacion para el fomento ensturias de la Investigacion Cientıfica Aplicada y la Tecnologıa”

FICYT) for the grant to Natalia Fidalgo-Used and financialupport from the Project BQU-2003-04671 from the Ministerioe Ciencia y Tecnologıa, Madrid (Spain).

eferences

[1] F. Wania, D. Mackay, Environ. Sci. Technol. 30 (1996) 390A.[2] Internet website: http://www.epa.gov/.[3] F.E. Ahmed, Trends Anal. Chem. 22 (2003) 170.[4] M. Petrovic, E. Eljarrat, M.J. Lopez de Alda, D. Barcelo, J. Chromatogr.

A 974 (2002) 23.[5] E. Bjorklund, T. Nilsson, S. Boward, Trends Anal. Chem. 19 (2000) 434.[6] C.A. de Wit, M. Alaee, D.C.G. Muir, Chemosphere 64 (2006) 209.[7] J.L. Domingo, J. Chromatogr. A 1054 (2004) 327.[8] A. Covaci, S. Voorspoels, L. Ramos, H. Neels, R. Blust, J. Chromatogr.

A (2006) (review available online).
[9] F.J. Santos, M.T. Galceran, Trends Anal. Chem. 21 (2002) 672.
[10] D. Barcelo (Ed.), Environmental Analysis, Techniques, Applications andQuality Assurance, Elsevier, Amsterdam, 1993.

[11] R.E. Majors, LC–GC Int. 4 (1991) 10.[12] R.M. Smith, J. Chromatogr. A 1000 (2003) 3.

himica Acta 590 (2007) 1–16

[13] V. Lopez-Avila, Crit. Rev. Anal. Chem. 29 (1999) 195.[14] V. Camel, Analyst 126 (2001) 1182.[15] J.R. Dean, G. Xiong, Trends Anal. Chem. 19 (2000) 553.[16] M.D. Luque de Castro, L.E. Garcia-Ayuso, Anal. Chim. Acta 369 (1998)

1.[17] K. Pointet, A. Milliet, Chemosphere 40 (2000) 293.[18] P.F. Hoekstraa, T.M. O’Harac, S.M. Backusa, C. Hannsc, D.C.G. Muira,

Environ. Res. 98 (2005) 329.[19] G. Asmun, K. Vorkamp, S. Backus, M. Comba, Sci. Total Environ. 331

(2004) 233.[20] S. Tanabe, M. Watanabe, T.B. Minh, T. Kunisue, S. Nakanishi, H. Ono,

H. Tanaka, Environ. Sci. Technol. 38 (2004) 403.[21] R.A. Hites, J.A. Foran, S.J. Schwager, B.A. Knuth, M.C. Hamilton, D.O.

Carpenter, Environ. Sci. Technol. 38 (2004) 4945.[22] I. Vives, J.O. Grimalt, J. Chromatogr. B 768 (2002) 247.[23] S. Corsolini, T. Romeo, N. Ademollo, S. Greco, S. Focardia, Microchem.

J. 73 (2002) 187.[24] J. Malavia, F.J. Santos, M.T. Galceran, J. Chromatogr. A 1056 (2004)

171.[25] S. Corsolini, K. Kannan, T. Imagawa, S. Focardi, J.P. Geasy, Environ. Sci.

Technol. 36 (2002) 3490.[26] K. Kannan, K. Hilscherova, T. Imagawa, N. Yamashita, L.L. Williams,

J.P. Giesy, Environ. Sci. Technol. 35 (2001) 441.[27] J.-H. Peng, C.-W. Huang, Y.-M. Weng, H.-K. Yak, Chemosphere 66

(2007) 1990.[28] I. Vives, J.O. Grimalt, S. Lacorte, M. Guillamon, D. Barcelo, Environ.

Sci. Technol. 38 (2004) 2338.[29] A.C. Gama, P. Sanatcumar, P. Viana, D. Barcelo, J.C. Bordado, Chemo-

sphere 64 (2006) 306.[30] K. Kannan, J.L. Reiner, S.H. Yun, E.E. Perrotta, L. Tao, B. Johnson-

Restrepo, B.D. Rodan, Chemosphere 61 (2005) 693.[31] P. Manirakiza, A. Covaci, L. Nizigiymana, G. Ntzakimazi, P. Schepens,

Environ. Pollut. 117 (2002) 447.[32] J. de Boer, C. Allchin, R. Law, B. Zegers, J.P. Boon, Trends Anal. Chem.

20 (2001) 591.[33] J.H. Christensen, M. Glasius, M. Pecseli, J. Platz, G. Pritzl, Chemosphere

47 (2002) 631.[34] C.R. Allchin, R.J. Law, S. Morris, Environ. Pollut. 105 (1999)

197.[35] R.J. Law, C.R. Allchin, M.E. Bennett, S. Morris, E. Rogan, Chemosphere

46 (2002) 673.[36] K. Vorkamp, J.H. Christensen, M. Glasius, F.F. Riget, Mar. Pollut. Bull.

48 (2004) 111.[37] C.-T. Fu, S.-C. Wu, Mar. Pollut Bull. 51 (2005) 932.[38] N.G. Dodder, B. Strandberg, T. Augspurger, R.A. Hites, Sci. Total Envi-

ron. 311 (2003) 81.[39] N. Fidalgo-Used, G. Centineo, E. Blanco-Gonzalez, A. Sanz-Medel, J.

Chromatogr. A 1017 (2003) 35.[40] S. Morris, P. Bersuder, C.R. Allchin, B. Zegers, J.P. Boon, P.E.G.

Leonards, J. de Boer, Trends Anal. Chem. 25 (2006) 343.[41] N. Holmqvist, P. Stenroth, O. Berglund, P. Nystrom, K. Olsson, D.

Jellyman, A.R. McIntosh, P. Larsson, Environ. Pollut. 141 (2006)532.

[42] K. Ramu, N. Kajiwara, S. Tanabe, P.K.S. Lam, T.A. Jefferson, Mar. Pollut.Bull. 51 (2005) 669.

[43] H. Kiviranta, T. Vartiainen, R. Parmanne, A. Hallikainen, J. Koistinen,Chemosphere 50 (2003) 1201.

[44] N. Hanari, Y. Horii, T. Okazawa, J. Falandysz, I. Bochentin, A.Orlikowska, T. Puzyn, B. Wyrzykowskaa, N. Yamashita, J. Environ.Monit. 6 (2004) 305.

[45] J. Pan, Y.-L. Yang, Q. Xu, D.-Z. Chen, D.-L. Xi, Chemosphere 66 (2007)1971.

[46] P. Isosaari, A. Hallikainen, H. Kiviranta, P.J. Vuorinen, R. Parmanne, J.
Koistinen, T. Vartiainen, Environ. Pollut. 141 (2006) 213.
[47] F. Priego-Capote, M.D. Luque de Castro, Trends Anal. Chem. 23 (2004)644.

[48] P. Rodrıguez-Sanmartın, A. Moreda-Pineiro, A. Bermejo-Barrera, P.Bermejo-Barrera, Talanta 66 (2005) 683.

http://www.epa.gov/

tica C
[49] N. Ratola, S. Lacorte, A. Alves, D. Barcelo, J. Chromatogr. A 1114 (2006)198.

[50] M. Jin, Y. Zhu, J. Chromatogr. A 1118 (2006) 111.[51] K.E.C. Smith, G.L. Northcott, K.C. Jones, J. Chromatogr. A 1116 (2006)

20.[52] C. Domeno, M. Blasco, C. Sanchez, C. Nerın, Anal. Chim. Acta 569

(2006) 103.[53] Y.C. Ling, H.C. Teng, C. Cartwright, J. Chromatogr. A 835 (1999) 145.[54] V.G. Zuin, J.H. Yariwake, C. Bicchi, J. Chromatogr. A 985 (2003) 159.[55] C. Quan, S. Li, S. Tian, H. Xu, A. Lin, L. Gu, J. Supercrit. Fluids 31

(2004) 149.[56] X.R. Zhu, H.K. Lee, J. Chromatogr. A 976 (2002) 393.[57] M.O.P. Crespo, M.A.L. Yusty, Chemosphere 59 (2005) 1407.[58] B. van Bavel, E. Sundelin, J. LillbaIck, M. Dam, G. LindstroIm,

Organohal. Compd. 40 (1999) 359.[59] H. Wolkers, M.O. Hammill, B. van Bavel, Environ. Pollut. 142 (2006)

476.[60] C. Nerın, R. Batlle, M. Sartaguda, C. Pedrocchi, Anal. Chim. Acta 464

(2002) 303.[61] P. Antunes, O. Gil, M.G. Bernardo-Gil, J. Supercrit. Fluids 25 (2003) 135.[62] V. Camel, Trends Anal. Chem. 19 (2000) 229.[63] C.S. Eskilsson, E. Bjorklund, J. Chromatogr. A 902 (2000) 227.[64] L. Cai, J. Xing, L. Dong, C. Wu, J. Chromatogr. A 1015 (2003) 11.[65] M. Barriada-Pereira, E. Concha-Grana, M.J. Gonzalez-Castro, S.

Muniategui-Lorenzo, P. Lopez-Mahıa, D. Prada-Rodrıguez, E.Fernandez-Fernandez, J. Chromatogr. A 1008 (2003) 115.

[66] M. Barriada-Pereira, M.J. Gonzalez-Castro, S. Muniategui-Lorenzo, P.Lopez-Mahıa, D. Prada-Rodrıguez, E. Fernandez-Fernandez, J. Chro-matogr. A 1061 (2004) 133.

[67] M. Barriada-Pereira, M.J. Gonzalez-Castro, S. Muniategui-Lorenzo, P.Lopez-Mahıa, D. Prada-Rodrıguez, E. Fernandez-Fernandez, Chemo-sphere 58 (2005) 1571.

[68] N. Carro, I. Garcia, M. Llompart, Analusis 28 (2000) 720.[69] G. Wittmann, T. Huybrechts, H. Van Langenhove, J. Dewulf, H. Nollet,

J. Chromatogr. A 993 (2003) 71.[70] S. Bayen, H.K. Lee, J.P. Obbard, J. Chromatogr. A 1035 (2004) 291.[71] S. Bayen, O. Wurl, S. Karuppiah, N. Sivasothi, H.K. Lee, J.P. Obbard,

Chemosphere 61 (2005) 303.[72] T. Pena, L. Pensado, C. Casais, C. Mejuto, R. Phan-Tan-Luu, R. Cela, J.

Chromatogr. A 1121 (2) (2006) 163.[73] B.E. Richter, B.A. Jones, J.L. Ezzell, N.L. Porter, N. Avdalovic, C. Pohl,

Anal. Chem. 68 (1996) 1033.[74] L.J. Fitzpatrick, O. Zuloaga, N. Etxebarria, J.R. Dean, Rev. Anal. Chem.

19 (2000) 75.[75] H. Bautz, J. Polzer, L. Stieglitz, J. Chromatogr. A 815 (1998) 231.[76] L. Ramos, E.M. Kristenson, U.A. Th. Brinkman, J. Chromatogr. A 975

(2002) 3.[77] M. Weichbrodt, W. Vetter, B. Luckas, J. AOAC Int. 83 (2000) 1334.[78] P. Suchan, J. Pulkrabova, J. Hajslova, V. Kocourek, Anal. Chim. Acta 520

(2004) 193.[79] S. Tao, L.Q. Guo, X.J. Wang, W.X. Liu, T.Z. Ju, R. Dawson, J. Cao, F.L.

Xu, B.G. Li, Sci. Total Environ. 320 (2004) 1.[80] J.L.F. Moreno, F.J.A. Liebanas, A.G. Frenich, J.L.M. Vidal, J. Chro-

matogr. A 1111 (2006) 97.[81] K. Adou, W.R. Bontoyan, P.J. Sweeney, J. Agric. Food Chem. 49 (2001)

4153.[82] J. Haib, I. Hofer, J.-M. Renaud, J. Chromatogr. A 1020 (2003) 173.[83] J.-F. Focant, G. Eppe, C. Pirard, E. De Pauw, J. Chromatogr. A 925 (2001)

207.[84] E. Bjorklund, A. Muller, C. von Hols, Anal. Chem. 73 (2001) 4050.[85] J.L. Gomez-Ariza, M. Bujalance, I. Giraldez, A. Velasco, E. Morales, J.

Chromatogr. A 946 (2002) 209.[86] K. Kitamura, Y. Takazawa, S. Hashimoto, J.-W. Choi, H. Ito, M. Morita,

Anal. Chim. Acta 512 (2004) 27.[87] E. Martinez, M. Gros, S. Lacorte, D. Barcelo, J. Chromatogr. A 1047

(2004) 181.[88] M. Janska, M. Tomaniova, J. Hajslova, V. Kocourek, Anal. Chim. Acta

520 (2004) 93.

himica Acta 590 (2007) 1–16 15

[89] L. Liguori, K. Heggstad, H.T. Hove, K. Julshamn, Anal. Chim. Acta573–574 (2006) 181.

[90] I. Johansson, K. Heas-Moisan, N. Guiot, C. Munschy, J. Tronczynski,Chemosphere 64 (2006) 296.

[91] H.M. Stapleton, N.G. Dodder, J.R. Kucklick, C.M. Reddy, M.M. Schantz,P.R. Becker, F. Gulland, B.J. Porter, S.A. Wise, Mar. Pollut. Bull. 52(2006) 522.

[92] K. Saito, A. Sjodin, C.D. Sandau, M.D. Davis, H. Nakazawa, Y. Matsuki,D.G. Patterson Jr., Chemosphere 57 (2004) 373.

[93] Q. Jiang, N. Hanari, Y. Miyake, T. Okazawa, R.K.F. Lau, K. Chen, B.Wyrzykowska, M.K. So, N. Yamashitab, P.K.S. Lam, Environ. Pollut.(2006) (article available online).

[94] R. Carabias-Martınez, E. Rodrıguez-Gonzalo, P. Revilla-Ruiz, J.Hernandez-Mendez, J. Chromatogr. A 1089 (2005) 1.

[95] S. Morales-Munoz, J.L. Luque-Garcıa, M.D. Luque de Castro, Anal.Chem. 74 (2002) 4213.

[96] L. Wennrich, P. Popp, J. Breuste, Chromatographia 53 (2001) S380.[97] S.A. Barker, A.R. Long, C.R. Short, J. Chromatogr. 475 (1989) 353.[98] E.M. Kristenson, L. Ramos, U.A. Th. Brinkman, Trends Anal. Chem. 25

(2006) 96.[99] L. Ramos, J.J. Ramos, U.A. Th. Brinkman, Anal. Bioanal. Chem. 381

(2005) 119.[100] J.-J. Ramos, M.-J. Gonzalez, L. Ramos, J. Sep. Sci. 27 (2004) 595.[101] L. Pensado, M.C. Casais, M.C. Mejuto, R. Cela, J. Chromatogr. A 1077

(2005) 103.[102] M.R. Criado, D.H. Fernandez, I.R. Pereiro, R.R.C. Torrijos, J. Chro-

matogr. A 1056 (2004) 187.[103] A. Martınez, M. Ramil, R. Montes, D. Hernanz, E. Rubı, I. Rodrıguez,

R.C. Torrijos, J. Chromatogr. A 1072 (2005) 83.[104] A.M. Carro, R.A. Lorenzo, F. Fernandez, R. Rodil, R. Cela, J. Chromatogr.

A 1071 (2005) 93.[105] M.C.R. Camargo, M.C.F. Toledo, Food Control 14 (2003) 49.[106] M.K. So, X. Zhang, J.P. Giesy, C.N. Fung, H.W. Fong, J. Zheng, M.J.

Kramer, H. Yoo, P.K.S. Lam, Mar. Pollut. Bull. 51 (2005) 677.[107] S. Jarayaman, R.J. Pruell, R. Mckinney, Chemosphere 44 (2001) 181.[108] R. Serrano, F.J. Lopez, F. Hernandez, J. Chromatogr. A 855 (1999) 633.[109] I. Vives, J.O. Grimalt, P. Fernandez, B. Rosseland, Sci. Total Environ.

324 (2004) 67.[110] S. Yenisoy-Karakas, Anal. Chim. Acta 571 (2006) 298.[111] R.C. Prados-Rosales, J.L.L. Garcıa, M.D.L. de Castro, J. Chromatogr. A

993 (2003) 121.[112] K. Gustavson, P. Jonsson, Mar. Pollut. Bull. 38 (1999) 723.[113] M.M. Storelli, A. Storelli, R. D’Addabbo, G. Barone, G.O. Marcotrigiano,

Environ. Int. 30 (2004) 343.[114] G.A. Stern, C.R. Macdonald, D. Armstrong, B. Dunn, C. Fuchs, L. Har-

wood, D.C.G. Muir, B. Rosenberg, Sci. Total Environ. 351–352 (2005)344.

[115] A. Meudec, J. Dussauze, M. Jourdin, E. Deslandes, N. Poupart, J. Chro-matogr. A 1108 (2006) 240.

[116] A. Binelli, A. Provini, Ecotoxicol. Environ. Saf. 58 (2004) 139.[117] R. Fuoco, S. Giannarelli, M. Onor, A. Ceccarini, V. Carli, Microchem. J.

79 (2005) 69.[118] S. Sinkkonen, J. Paasivirta, Chemosphere 40 (2000) 619.[119] G. Xiong, X. He, Z. Zhang, Anal. Chim. Acta 413 (2000) 49.[120] G. LindstoIm, H. Wingfors, M. Dam, B. van Bavel, Arch. Environ. Con-

tam. Toxicol. 36 (1999) 355.[121] M. Zennegg, M. Kohler, A.C. Gerecke, P. Schmid, Chemosphere 51

(2003) 545.[122] B. Gomara, C. Garcıa-Ruiz, M.J. Gonzalez, M.L. Marina, Anal. Chim.

Acta 565 (2006) 208.[123] K. Vorkamp, J.H. Christensen, M. Glasius, F.F. Riget, Marine Pollut. Bull.

48 (2004) 111.[124] F. Riget, K. Vorkamp, R. Dietz, S.C. Rastogi, J. Environ. Monit. 8 (2006)

1000.[125] K.S. Kumar, K. Kannan, J.P. Giesy, S. Masunaga, Environ. Sci. Technol.

36 (2002) 2789.[126] J.-F. Focant, C. Pirard, E. De Pauw, Talanta 63 (2004) 1101.[127] C. Pirard, E. De Pauw, J.-F. Focant, J. Chromatogr. A 998 (2003) 169.

1 tica C

[152] M. Jaremo, E. Bjorklund, N. Nilsson, L. Karlsson, L. Mathiasson, J.


[128] J. Zrostokova, S.J. Lehotay, J. Hajslova, J. Sep. Sci. 25 (2002) 527.[129] Z.A. Kuzyk, J.P. Stow, N.M. Burgess, S.M. Solomon, K.J. Reimer, Sci.

Total Environ. 351–352 (2005) 264.[130] M.J. Ryan, G.A. Stern, M. Diamond, M.V. Croft, P. Roach, K. Kidd, Sci.

Total Environ. 351–352 (2005) 501.[131] M.D.L. Easton, D. Luszniak, E. Von der Geest, Chemosphere 46 (2002)

1053.[132] R.J. Norstrom, M. Simon, J. Moisey, B. Wakeford, D.V.C. Weseloh,

Environ. Sci. Technol. 36 (2002) 4783.[133] A. Olsson, M. Vitinsh, M. Plikshs, A. Bergman, Sci. Total Environ. 239

(1999) 19.[134] W. Vetter, M. Weichbrodt, E. Scholz, B. Luckas, H. Oelschlager, Mar.

Pollut. Bull. 38 (1999) 830.[135] J. Verreault, D.C.G. Muir, R.J. Norstrom, I. Stirling, A.T. Fisk, G.W.

Gabrielsen, A.E. Derocher, T.J. Evans, R. Dietz, C. Sonne, G.M. Sandala,W. Gebbink, F.F. Riget, E.W. Born, M.K. Taylor, J. Nagy, R.J. Letcher,Sci. Total Environ 351–352 (2005) 369.

[136] A. Evenset, G.N. Christensen, T. Skotvold, E. Fjeld, M. Schlabach, E.Wartena, D. Gregor, Sci. Total Environ. 318 (2004) 125.

[137] J.M. Luross, M. Alaee, D.B. Sergeant, C.M. Cannon, D.M. Whittle, K.R.
Solomon, D.C.G. Muir, Chemosphere 46 (2002) 665.
[138] K. Patel, R.J. Fussell, M. Hetmanski, D.M. Goodall, B.J. Keely, J. Chro-matogr. A 1068 (2005) 289.

[139] D.P. Herman, J.I. Effler, D.T. Boyd, M.M. Krahn, Mar. Environ. Res. 52(2001) 127.

himica Acta 590 (2007) 1–16

[140] H. Lord, J. Pawliszyn, J. Chromatogr. A 885 (2000) 153.[141] C.L. Arthur, J. Pawliszyn, Anal. Chem. 62 (1990) 2145.[142] H. Kataoka, H.L. Lord, J. Pawliszyn, J. Chromatogr. A 880 (2000)

35.[143] A.A. Boyd-Boland, S. Madgic, J. Pawliszyn, Analyst 121 (1996)

929.[144] G. Theodoridis, E.H.M. Koster, G.J. de Jong, J. Chromatogr. B 745 (2000)

49.[145] M. de Fatima Alpendurada, J. Chromatogr. A 889 (2000) 3.[146] J. Beltran, F.J. Lopez, F. Hernandez, J. Chromatogr A 885 (2000) 389.[147] A. Penalver, E. Pocurull, F. Borrull, R.M. Marce, Trends Anal. Chem. 18

(1999) 557.[148] D.A. Lambropoulou, I.K. Konstantinou, T.A. Albanis, J. Chromatogr. A

1124 (2006) 97.[149] D.A. Lambropoulou, I.K. Konstantinou, T.A. Albanis, Anal. Chim. Acta

573–574 (2006) 223.[150] W.-H. Ho, S.-J. Hsieh, Anal. Chim. Acta 428 (2001) 111.[151] R. Rodil, A.M. Carro, R.A. Lorenzo, R. Cela Torrijos, Anal. Chem. 77

(2005) 2259.

Chromatogr. A 877 (2000) 167.[153] S. Sporring, E. Bjorklund, J. Chromatogr. A 1040 (2004) 155.[154] E. Eljarrat, A. de la Cal, D. Raldua, C. Duran, D. Barcelo, Environ. Sci.

Technol. 38 (2004) 2603.

sample handling strategies for the determination of persistent trace organic contaminants from biota...

Documents

sample extraction

extraction techniques

analytes extraction

extraction mae

rated extraction

sample preparation steps

determination of analytes

determination of pops