Applications of solid phase microextraction for ... - Wiley Online Library

J. Sep. Sci. 2006, 29, 333 – 345

Varinder Kaur1 Ashok Kumar Malik1 Neelam Verma2 1

Department of Chemistry, Punjabi University, Patiala, Punjab, India 2 Department of Biotechnology, Punjabi University, Patiala, Punjab, India

V. Kaur et al.

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Review Article Applications of solid phase microextraction for the determination of metallic and organometallic species This paper reviews recent developments of solid phase microextraction (SPME) and its application to the analysis of organometallic species of lead, arsenic, mercury, tin, and selenium by hyphenation with HPLC-GC-atomic spectrometry. In the first part, a background of the technique is given in terms of derivatization, fibers used, extraction and desorption conditions. The second part summarizes typical SPME applications to the determination of organometallic species and the main experimental conditions with the aid of specific examples. Most of the applications comprise alkylation with NaBEt4 and headspace extraction followed by gas chromatographic separation with a suitable detector. Keywords: GC / HPLC / Metallic and organometallic species / Solid phase microextraction / Received: August 24, 2005; revised: November 10, 2005; accepted: November 10, 2005 DOI 10.1002/jssc.200500319

1 Introduction The commercial use of organometallic compounds has increased in recent decades, leading to greater direct interaction of organometals with the environment. Currently, the most important and abundant organometallic species in the environment are organomercury, orga-

Correspondence: Dr. Ashok Kumar Malik, Department of Chemistry, Punjabi University, Patiala-147 002, Punjab, India. E-mail: [email protected] Fax: +91-175-2283073 Abbreviations: AAS, atomic absorption spectrophotometry; AED, atomic emission detector; AES, atomic emission spectrophotometry; AsB, arsenobetaine; BT, butyltin; CAR, carboxen; CGC, capillary gas chromatography; CW, carbowax; DBDET, dibutyldiethyltin; DBT, dibutyltin; DEM, diethylmercury; DES, diethyl selenide; DI, direct immersion; DMA, dimethylarsonic acid; DMDS, dimethyl diethyl selenide; DMM, dimethylmercury; DMS, dimethyl selenide; DMT, dimethyltin; DPM, diphenylmercury; DVB, divinylbenzene; EA-SPME, electrochemically aided solid phase microextraction; EC, electrochemical; ES, electrospray; FID, flame ionization detector; HS, headspace; HT, hexyltin; ICP-MS, integrated coupled plasma mass spectrometry; LLE, liquid liquid extraction; MBT, monobutyltin; MC-GC, multicapillary gas chromatography; MEM, monoethylmercury; MIP, microwave induced plasma; MMM, monomethylmercury; MMT, monomethyltin; MOT, monooctyltin; MPM, monophenylmercury; MT, methyltin; PA, polyacrylate; PDMS, polydimethylsiloxane; PED, plasma emission detector; PFPD, pulsed flame photometry; PT, phenyltin; SPME, solid phase microextraction; TBET, tributylethyltin; TBT, tributyltin; TeBT, tetrabutyltin; TEL, tetraethyllead; TML, trimethyllead; TMPT, trimethylphenyltin; TMT, trimethyltin; TOFMS, time of flight mass spectrometry; TPA, triphenylarsine; TPT, triphenyltin.

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2006 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim

nolead, and organotin compounds. These compounds cause serious environmental or toxicity problems [1]. For the speciation of organometallic compounds, various hyphenated chromatographic techniques are used which are coupled with a suitable detector. Mostly, gas chromatography is used for the speciation of thermally stable and volatile species and the method generally involves alkylation of the analyte prior to its extraction. Those species which are not thermally stable or are nonvolatile can be easily separated by HPLC. The extensive use of organic solvents is no longer desirable as these are expensive and harmful to the environment and to health. A number of methods have been developed which are solvent free or low solvent consumption methods. Among these, SPE [2] reduces the limitations of liquid-liquid extraction (LLE) but it is still time consuming. In recent years, SPME [3, 4] has been proposed as a promising alternative to LLE due to its simplicity of use, high preconcentration power, and ability to extract volatile methylated species as well as phenyl derivatives. It is used in hyphenation with GC and HPLC for the determination of various organometallic species [5]. Detection methods such as integrated coupled plasma mass spectrometry (ICP-TOFMS), microwave induced plasma-atomic emission spectrometry (MIPAES), atomic emission detection (AED), flame ionization detection (FID), pulsed flame photometric detection (PFPD), and atomic absorption spectrophotometry (AAS) are used in the case of GC for such determinations. Reviews of the determination of organometallic species www.jss-journal.com

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(organotin and organomercury) are given in the literature [6 – 8]. This review focuses primarily on all SPME methods for the analysis of metallic and organometallic species of Sn, Pb, Hg, As, and Se.

2 Derivatization Ionic organometallic compounds require derivatization prior to extraction. Derivatization is a process of chemically modifying a species to produce a new species which has properties more suitable for analysis. A number of derivatizing methods have been employed to coordinate the metal ions [9, 10]. The main advantages of derivatization are: (i) it increases the volatility of the species; (ii) it increases the stability of the species; (iii) it improves the chromatographic behavior or detectability of the complex and (iv) it enhances sensitivity. Most of the conventional SPME coatings are not compatible with an organic-solvent-rich environment, so the derivatizing step is carried out directly in the aqueous phase. This restricts the choice of derivatizing agent to the alkylborates as these reagents can tolerate aqueous conditions [11]. The use of tetraalkylborates such as NaBEt4, and NaBPh4 for the determination of organometallic species has been reported in the literature. Most derivatizing agents tend to be moisture- and light sensitive, and thus require special handling [12]. Recent investigations with various dry aprotic solvents such as THF have shown that reagent stability can be improved by storage under cool conditions [13]. The use of alkyl- or arylborates as derivatizing reagent has limitations in the simultaneous determination of ethyl, butyl, and phenyl species, especially environmentally relevant butyltin, phenyltin, and ethyllead species, without the loss of species information [14]. As an example, the derivatization of lead species is critical. It is possible to use sodium tetrahydroborate (NaBH4) and sodium tetraethylborate to derivatize the lead species. However, in the case of sodium tetrahydroborate, the hydrides generated tend to dismutate due to their poor stability. The other reagent, sodium tetraethylborate can be used to derivatize methyllead species, but full speciation of ethyllead is not possible as both ethyllead and inorganic lead compounds yield Pb(C2H5)4 and eliminate species-specific information. To overcome this problem, tetra(n-propylborate) [14] and tetraammonium tetrabutylborate [15] are used to derivatize the organometals in the aqueous samples. However, in the case of lead species, extraction by organic solvent is required after derivatization. To remove the inorganic lead, addition of a chelation reagent is also needed. Thus, determination of organolead and inorganic lead is not possible with these reagents. Deuterated sodium ethylborates [16] are therefore used for the derivatization of lead species. Other

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methods of derivatization involve hydride generation [17, 18] and Grignard derivatization. Grignard derivatization [10, 19, 20] requires aprotic media and, therefore, numerous handling steps. It is a complicated process and involves consumption of both reagent and time [21]. Sometimes, derivatization causes problems in GC determination due to the lower stability of non-ionic alkyllead compounds, so ionic organolead compounds can be determined directly by using electrospray MS (ES-MS). [22, 23]. In such cases, in-tube SPME is used for preconcentration in which the extraction phase coats the inside of an open tube through which the sample is passed. Analytes in aqueous samples are directly extracted and concentrated into the stationary phase of the capillary column by repeated draw/eject cycles of sample solution, and they can be directly transferred to the liquid chromatographic system. The in-tube SPME technique saves preparation time, solvent purchase and disposal cost, and can improve detection limits. On line in-tube SPME can automatically perform continuous extraction, concentration, desorption, and injection using an autosampler, and is generally employed in combination with HPLC and LC-MS. Derivatization can improve efficiency, selectivity, and subsequent GC detection. Derivatization can be accomplished directly on the SPME fiber and in the GC injector port. In direct derivatization, the reagent is added directly to the sample. The derivatized analytes can be extracted onto the SPME fiber and analyzed by GC with a suitable detector. In injector port derivatization, a fiber is placed in the aqueous phase to isolate the analytes. After the desired extraction time, the fiber is transferred into the hot GC injector port for desorption, derivatization, separation, and quantification. Sodium tetraethylborate is used [24, 25] for the determination of organometallic species.

3 SPME fiber Analytes can be extracted from a matrix by using an SPME fiber. In this technique, analytes are partitioned between the matrix and a stationary phase coated on the fiber. The amount of analyte extracted depends upon the partition coefficient of the analyte. Seven different types of fibers are commercially available [Supelco, Bellefonte, PA, USA]. Selection of the type of polymer used for the extraction depends upon the chemical nature of the analyte, i. e., polarity and volatility. In general, polar fibers are used for polar analytes and non-polar ones for nonpolar analytes. Polydimethylsiloxane is the most useful liquid type coating. It has been used in various applications of SPME to the analysis of organometallic species (see Table 1). However, some other adsorptive coatings have been used for the determination of organometals www.jss-journal.com

J. Sep. Sci. 2006, 29, 333 – 345

The coating should be thin to reduce the extraction times because thick coatings require more time to equilibrate [30]. Coatings can be attached to the fused-silica core by various methods. Commercially available fibers can be damaged in strong organic solvents, strong acids, and alkali solutions. To overcome this problem, Chong et al. developed sol-gel coatings [31]. Gbatu et al. [32] used fibers prepared by sol-gel technology which are stable even in strong organic solvents (xylene and methylene chloride) as well as in acidic and basic solution. The preparation of sol-gel fibers involves four steps: (i) pretreatment of the fused-silica fiber to remove the polyimide coating. This exposes a maximum number of silanol groups. It can be achieved by burning the tip of the fiber; (ii) washing and drying of the fiber; (iii) exposure of the fiber tip to the so-gel solution for 20 min and (iv) conditioning of the fiber placing it in the GC injector port at a temperature of 1308C. Because the sol-gel coating is chemically bonded to the surface of the fused-silica fiber, these fibers are more stable than commercially available fibers. Figure 1 shows a schematic diagram of a sol-gel SPME fiber. Sol-gel PDMS fibers can be used up to 3208C whereas commercial PDMS fibers begin to bleed at lower temperatures (2008C). The high porosity of sol-gel fibers results in higher sensitivity and faster extraction times than for commercial fibers.

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vapor phase above the sample. In this case, the fiber is not in direct contact with the sample. The analytes need to be transported through a layer of air before they can reach the coating. In DI-SPME the coated fiber is immersed in the liquid sample and the analytes are transported directly to the extraction phase. For volatile organometallic compounds, fiber HS-SPME is preferred over DI-SPME [30]. Recently SPME practitioners have extended the approach to speciation of monomethylarsonic acid and dimethylarsinic acid by using DI-SPME [33]. Extraction efficiency can be improved by modification of matrix, target analytes, and fiber chemistry. It is reported in the literature that the extraction yield for DVB/CAR/PDMS fibers is much higher than that observed with PDMS fibers in the extraction of organometallic compounds of mercury, lead, and tin [29]. Extraction of the analyte in the case of in-tube SPME involves the washing of the capillary with the sample solution by applying draw from the sample-eject into the sample’ extraction cycles. The cycles are repeated until equilibrium is reached. The valve is then switched to the inject position and mobile phase passes through the extraction capillary. ,

such as carboxen/polydimethylsiloxane (CAR/PDMS) [26, 27], polydimethylsiloxane/divinylbenzene (PDMS/DVB) [26, 28], divinylbenzene/carboxen/polydimethylsiloxane (DVB/CAR/PDMS) [29], carbowax/polydimethylsiloxane (CW/PDMS) [24], and polyacrylate (PA) [28].

Solid phase microextraction

5 Desorption Analysis involves desorption of the organometallic compounds into the desorption chamber of a GC or an HPLC instrument. The two methods have different desorption steps. In most applications, SPME is hyphenated to GC, but recent studies have reported the hyphenation of SPME to HPLC for the speciation of various analytes [34].

5.1 Desorption in GC

Figure 1. Schematic diagram of sol-gel SPME fiber assembly. (A) coated silica tip, (B) stainless steel piercing needle, (C) fused silica, (D) seal, (E) plain attachment hub. Reproduced with permission from [32].

4 Extraction and desorption conditions The fiber is exposed to the sample for a particular time period to extract the analyte. When equilibrium is established, the fiber is removed from the sample. Equilibration can be enhanced by using a magnetic stirrer, ultrasonication, or an elliptical shaker. The nature of the analyte, the thickness of the fiber, and the chemistry of the fiber greatly affect the time required for the equilibration. There are two modes of fiber SPME for extraction of analytes: Headspace (HS) SPME and direct immersion (DI) SPME. HS-SPME involves exposure of the fiber to the

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In this case analytes are thermally desorbed into the injector of a GC. The method is used for volatile and thermally stable compounds. After extraction of analyte, the fiber is introduced into the injector port to effect thermal desorption of analytes from the coating.

5.2 Desorption in HPLC In the case of HPLC, desorption can be performed in a specially designed six port injector with a desorption chamber installed in place of a sample loop. Organic solvent is used as a mobile phase for the desorption of analytes. This approach has distinct advantages because thermal desorption at high temperature in GC leads to incomplete desorption of non-volatile compounds and to degradation of the polymer coating. Two modes of desorption are used in HPLC, viz. dynamic and static desorption mode. The dynamic mode of desorption is sufficient if the analyte is not strongly adsorbed onto the fiber. In this case, the analyte can be removed by a stream of mobile phase [32]. However, if the analytes are more www.jss-journal.com

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Table.1. Various parameters for the SPME of organometallic species. No.

Analyte

Matrix

Technique

Derivatizing agent

Fiber used, extraction time

Buffer

1

Organotin, organolead

Aqueous samples

SPME-HPLC-UV

NaBPr4 (in situ)

10 mm PDMS, magnetically stirred, 20 min

2

Organomercury, organolead, organotin (MBT, DBT, TBT) TEL,TML

Aqueous samples

SPME-CGC-ICPMS

NaBEt4 1%

–

In-tube SPME-HPLC-ES-MS

No derivatization

10 mm PDMS, magnetically stirred, 10 min In-tube GC capillary column

HS-SPME-GC-AED

NaBEt4 (in situ)

HS-SPME-MC-GCICP-TOFMS

–

0.1 M H2PO4– /H3PO4 0.2 ng/L (pH 3), acetic acid/ sodium acetate (pH 4 – 6), HPO24 – /H2PO4 – (pH 7) 0.2 mol/L acetic acid 0.34 – 2.1 ng/L and sodium acetate sol. (pH – 5.5) 0.1% TFA + 12% methanol as mobile phase, 0.45 mL/min (a) ammonia/citrate pg/L and ng/L buffer (pH 8.5) (b) sod. acetate buffer (pH 5) (a) below pg/g – (b) 2 pg/g (c) 1.3 pg/g

3

4

5

6

7

8 9 10

(a) Organotin, Aqueous (b) organolead samples (c) organoarsenic (d) organomercury, Biological (a) TML, DML, TMT, DMT, materials and MMT, MBT, DBT, TBT (b) 2+ road dust MMM (c) Hg Organomercury, organotin, Natural water 2+ organolead (Hg , MMM, TEL, TML, MBT, TBT)

Organotin, organomercury, Surface water organolead and sediment sample 2+ + + Pb , Pb(CH3)3, Pb(C2H5)3, Water samples Pb(C2H5)4 DMA, MMA Organotin, organolead 2+

Aqueous samples Water

HS-SPME-GC-EI-MS 2% NaBEt4

(a) 100 mm PDMS (better) (b) 75 mm CW/PDMS (a) PDMS/ DVB (b) and (c) CAR/PDMS (a) 100 mm PDMS 0.2 M acetic acid and (b) 30 mm DVB/CAR/ 0.2 M sodium acetate PDMS (better) (pH 5.5) magnetically stirred, 30 min PDMS fiber –

15

21

22, 23

24

26

Below ng/L or sub ng/L

29

0.13 – 3.7 ng/L

37

–

HS-SPME-GC-MS/ FID SPME-GC-ion trapMS SPME-GC-AED

2% deuterium 100 mm PDMS labeled NaB(C2D5)4 100 mm PDMS

Sodium acetate buffer (pH 4) –

95 ppt, 130 ppt 39 83 ppt, 90 ppt – 33

NaBEt4

–

–

0.09 pg, 0.08 pg 35

HS-SPME-GC-MS

–

100 mm PDMS, 10 min PDMS, 10 min Sol-gel fiber, magnetically stirred, (30 min, 25 min, 20 min) –

–

–

36

– 80 : 20% v/v acetonitrile/water, 1 mL/min

– 80, 412, 647 mg/L

38 32

TEL, Pb

12 13

Lead derivatives Organoarsenic, organomercury, organotin (TPA, DPM, TMPT)

Water Aqueous samples

HS-SPME-AAS SPME-HPLC-UV

– –

14

TEL, TML and DMA

Soil samples

–

15

Fish, waters

16

Methyl mercury derivatives AsB

HS-SPME-GCMIP-AED HS-SPME-GC-MS

17

AsB

18

Hg2+

Aqueous samples

SPME-ion trap-GC-MS

–

19

Hg2+ salts

SPME-GC-MS

–

20

MMM, MEM, MPM

HS-SPME-GC-AAS

–

21

DMM, DEM

SPME-GC-MIP-AED

22

24 25 26

Hg2+, organomercury compounds HgCH+3, Hg2+, MBT, DBT, TBT Methyltins DBDT, TBET, TeBT MBT, MPT

(a) Soils and water (b) slurry In soils from orchards and wheat samples Waters, soils, and environmental samples Seawater

100 mm PDMS, 10 min stirring Platinum wire with poly(3-dodecylthiophene) coating, (2 cm60.2 mm) Platinum wire with poly(3-alkylthiophene) coating (2 cm60.2 mm Carbon steel wire with 10 mm gold coating –

27

Organotin

i

Ref.

SPME-CGC-ICPMS

11

23

Detection limit

ICP-MS/SPMEHPLC-ICP-MS

NaBEt4 –

ICP-MS/SPMEHPLC-ICP-MS

–

40

–

–

41

30 mM (NH4)CO3, isocratic, flow rate 1 mL/min

–

42

Mobile phase – 30 mM (NH4)CO3, isocratic, flow rate 1 mL/min

43

–

–

44

–

(a) 7 ppb (b) 2 ppm

45

–

0.1 M sodium acetate buffer (pH 4)

16, 12, 7 ng

46

–

–

–

ng/g, mg/L

47

HS-SPME-GC-AES

NaBPh4

PDMS coated silica fiber

–

–

48

Aqueous samples Water River water Water

SPME-GC-MIP-AES

–

–

–

49

HS-SPME-GC-FPD SPME-GC-MS SPME-GC-FPD

NaBEt4 NaBEt4 2% NaBEt4

Lard fat

SPME-GC-MS

–


10 mm PDMS 100 mm PDMS 100 mm PDMS (best results), 65 mm PDMS-DVB, 65 mm CW-DVB, 85 mm PA

– – – 1 – 50 ppb Sodium ethanoate/etha- 2 – 4 ng/L noic acid (pH 4.8)

50 51 28

–

52

–

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Table.1. Continued ... No.

Analyte

Matrix

28

TPT

29

MBT, DBT, TBT

30

Butyltin

Aqueous stan- HS-SPME-CGC-ICPdard solution, MS potatoes, mussel Marine sediHS-SPME-GC-MS ments Environmental SPME-GC-FPD samples

31

(a) MBT (b) MPT

32

Organotins

33

Butyltin, phenyltin

34

Organotins

35

MBT, DBT,TBT

36

TPT, TBT

37

Organotins (MMT, MBT, MPT, MOT

38

Organotins

39

Organotins

40

BT, PT, HT

41

Organoselenium (DMS, DES, DMDS) Organoselenium (DMS, DES, DMDS)

42

Technique

Derivatizing agent

Fiber used, extraction time

Buffer

Detection limit

Ref.

NaBEt4

100 mm PDMS, 10 – 20 min

0.2 mol/L ammonium buffer (pH 8)

2 pg/L

53

NaBEt4 (in situ)

PDMS

–

730 – 969 pg/g

25

Grignard derivatization, hydrogenation NaBEt4

–

–

–

54

Natural SPME-GC-FPD aqueous samples, sediments, sewage sludge Fresh waters, SPME-GC-ICP-AES 1% NaBEt4 wastewater, sediments SPME-CGC-ICP-AES 1% NaBEt4

PDMS, 60 min

–

(a) 0.006 – 0.031 ng/L (b) 0.2 – 0.6 ng/L

55

100 mm PDMS, 15 min

–

–

56

100 mm PDMS, 40 min

Sediments, fish tissue Estuarine superficial sediment In French beans and potatoes, aqueous plants Spiked fish and waters

SPME-GC-FPD/PFPD/ – MIP-AES/ICP-MS HS-SPME-GC-FID NaBEt4

–

Sodium ethanoate/ ethanoic acid buffer (pH 4.8) –

100 mm PDMS, 15 min

Harbor sediments Seawater samples

SPME-GC-PFPD

0.2% NaBEt4

0.25 mm PDMS, 20 min

HS-SPME-GC-PFPD

1% NaBEt4

SPME-GC-PFPD

–

(a) CAR/PDMS (better) (b) PDMS –

HS-SPME-GC-MS

2% NaBEt4

Water

SPME-CGC

NaBEt4

Yeast

HS-SPME-MIP-AES

–

Yeast

HS-SPME-MC-GC– ICP-MS/MIP-AES/AFS

strongly adsorbed onto the fiber, it is dipped into the mobile phase or another strong solvent for a specified time. Desorption performed in this way is known as static desorption [22, 23]. Each type of desorption should be undertaken using a minimum quantity of solvent.

6 Applications of SPME Various applications of SPME to the determination of organometallic compounds are summarized in this part of the review. All pertinent parameters such as derivatization mode, fiber used, extraction time, buffer used, and detection limits are given in Table 1.

6.1 Organolead compounds Tutschku et al. [35] preconcentrated and determined organotin and organolead species in environmental samples by SPME and GC-AED. Analytical variables of the extraction such as adsorption and desorption time, stirring rate, and temperature were investigated. The proposed procedure has been successfully applied to the

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100 mm PDMS coated fused-silica fiber, 30 min 100 mm PDMS, 30 min – –

57

0.6 – 20 pg/L

58

1.5 M sodium acetate buffer (pH 4.3)

–

59

Sodium ethanoate/ ethanoic acid buffer (pH 4.8) Sodium acetate/acetic acid buffer (pH 4.8)

0.1 mg/kg

60

Sod. acetate/acetic acid buffer (pH 4.8) Acetate buffer (pH 5)

–

61

0.4 – 4.6 ng/L

62

–

–

63

–

Low ppb levels

64

–

0.057, 0.47, 0.19 ng/mL, respectively

65

0.02 and 56 ng/L 27

analysis of organotin compounds in various slurry samples. Gorecki et al. [36] determined tetraethyllead (TEL) and inorganic lead in water samples by SPME-GC. HS-SPME was used for extraction of inorganic lead after derivatization with sodium tetraethylborate to form TEL. The method was optimized with respect to pH, amount of derivatizing reagent added, stirring conditions, and extraction time. Centineo et al. [29] hyphenated GC with SPME by using mass spectrometry for detection of organometallic compounds of mercury, tin, and lead in natural waters (Fig. 2). The sample was derivatized with NaBEt4 and extracted by HS-SPME. The sample was separated and detected by GC-MS. All the important process parameters, such as SPME fiber coating, extraction time and extraction temperature, were optimized. Moens et al. [21] determined organomercury, organolead, and organotin compounds by HS-SPME-CGC-ICPMS. Sodium tetraethylborate was used for in-situ derivatizawww.jss-journal.com

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of ES-MS permits the simultaneous monitoring of both elemental and molecular forms of lead by applying different fragmentation voltages for elemental and molecular ions. The chromatogram obtained for TML and TEL by this method is shown in Fig. 4. Fragueiro et al. [38] determined TEL by SPME-thermal desorption-quartz-furnace AAS in gasoline (leaded and unleaded) and water. Three different volatilizers were designed and their influence on the thermal desorption of TEL was studied. HS extraction was done in stirred samples for 10 min. The sample was analyzed by AAS. Figure 2. HS-SPME-GC-MS chromatogram in SIM mode for the ethyl-derivative of a marina sample: 1) CH3Hg+; 2) TML; 3) Hg2+; 4) TML; 5) MBT; 6) DBT; 7) TBT. Reproduced with permission from [29].

tion. The derivatized complexes were sorbed on a PDMScoated fused-silica fiber, and desorbed in the splitless injection port of the GC. They were determined by ICPMS coupled to GC. De Smaele et al. [37] successfully determined organometallic compounds of Sn, Hg, and Pb in surface water and sediment samples by SPME–CGC–ICP-MS. The organometallic compounds were determined by in-situ derivatization in the aqueous phase. They were simultaneously extracted onto a PDMS fiber. This was followed by thermal desorption in the GC injection liner. HS-SPME favors sampling of species of interest and prevents interference by matrix components. Mester et al. [22, 23] extended the potential of SPME by hyphenating SPME and HPLC for the speciation of tetramethyllead (TML) and TEL. In this method, in-tube solidphase microextraction was hyphenated with ES-MS. The hyphenation of SPME–ES-MS is shown in Fig. 3. The use

Yu et al. [39] worked on the speciation of alkyllead and inorganic lead in water by using deuterium-labeled sodium tetraethylborate as in-situ derivatizing reagent. Derivatization was carried out directly in water and the derivatives were extracted by HS-SPME. The extracted analytes were then separated and detected by transfer to GC-MS or GC-FID. Different species such as Pb2+, Pb(CH3)3+, Pb(C2H5)3+, and Pb(C2H5)4 can be analyzed by this method (Fig. 5). This method was used to monitor the degradation of TEL in water. Mothes et al. [24] coupled SPME and GC-AED for the determination of organometallic compounds. To obtain a high extraction yield, the SPME conditions were optimized for each element by appropriate SPME fiber selection and variation of the exposure time, stirring rate, pH range, and desorption time. All the organometallic compounds tested were extracted from the aqueous phase using SPME. Crnoja et al. [14] reported a method for the simultaneous derivatization and SPME of organotin and organolead compounds in aqueous samples which makes use of GCAED. Sodium tetrapropylborate was used for derivatization. Use of the tetrapropylborate reagent allows the

Figure 3. Schematic of the in-tube SPME-RPHPLC-ES-MS system. Reproduced with permission from [22].

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Figure 4. Chromatogram of TML and TEL obtained by in-tube SPME-HPLC-ES-MS. The concentration of each compound is 50 ng/ mL. Both chromatograms were recorded simultaneously by monitoring the molecular masses (m/z 253, 295) of TML and TEL and, in parallel, the elemental lead mass (m/z 208). Reproduced with permission from [23].

Rosenkranz et al. [40] studied the behavior of different organometallic compounds in the presence of inorganic Hg2+. Transalkylation of mercury species and their analysis by the GC-MIP-AED system has been reported, in which four different Hg species are formed by the abiotic reaction of inorganic Hg with organolead and organoarsenic compounds. Transfer of one or two alkyl groups to the inorganic Hg is possible under the given conditions. Jitaru et al. [26] described an efficient method for the simultaneous speciation analysis of ten organometallic compounds of mercury, tin, and lead by SPME-MC-GC hyphenated to ICP-TOFMS. The sample was derivatized with NaBEt4 and extracted by HS-SPME. A 65-mm PDMS/ DVB fiber offered the best overall extraction efficiency among the seven different fibers used for comparing the extraction efficiency.

6.2 Organoarsenic compounds

Figure 5. (a) Total ion chromatogram of TML, DML, Pb2+, TEL, and TeEL after derivatization with NaBEt4. (b) Single ion chromatogram of Pb2+ derivative (Pb2+, m/z 309). (c) Single ion chromatogram of TEL derivative (TEL, m/z 299). (d) Single ion chromatogram of TeEL derivative (TeEL, m/z 294). Reproduced with permission from [39].

simultaneous derivatization and determination of all butyltin and phenyltin (BT and PT) and organolead (methyl- and ethyllead) compounds present in the environment.

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Cai et al. [41] determined methylmercury and labile Hg2+ in fish and river water matrices by using SPME. Derivatization of ionic mercury species was accomplished with sodium tetraethylborate in a sample vial and subsequent extraction with a PDMS-coated silica fiber. The analytes were desorbed in the GC and subsequently analyzed by EI-MS. Both HS-SPME and aqueous-phase SPME were studied. The method was applied to analyses of standard reference materials and river water samples. Gbatu et al. [32] widened the application of SPME by describing a method for the preparation of fibers which are highly stable in strong organic solvents (xylene and methylene chloride) as well as in acidic and basic solutions (pH 0.3 and 13). These fibers were used to extract organoarsenic, organomercury, and organotin compounds from aqueous solutions prior to separation using HPLC with UV absorbance detection. Different chromatograms are obtained by varying the ratios of C8-TEOS/ www.jss-journal.com

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external potential. This potential provides a sufficient driving force for desorption of the analyte from the extraction phase into an aqueous solution for subsequent analysis. The applied positive potential oxidizes the polymer to its charged hydrophilic state, which releases the neutral analyte. Quantitation and speciation of the analyte from the sample matrix were accomplished by HPLC coupled to ICP-MS. Tamer et al. [43] applied electrochemically aided SPME (EA-SPME) to the determination of neutral species such as arsenobetaine (AsB) by using electro-synthesized organic conducting polymer films. The separation and detection of the As species was attained using an HPLC-ICP-MS interfaced system. Comparison of the performance of poly(3-octylthiophene), poly(3-dodecylthiophene), and poly(3-hexadecylthiophene) films is reported. Determination of arsenic compounds in the presence of organometallic compounds of Sn, Pb, and Hg are also reported in [24, 40].

6.3 Organomercury compounds

Figure 6. Extraction of aqueous solution of 0.5 mg/L Ph3As, 5 mg/L Ph2Hg, and 25 mg/L TMPT with fibers prepared from sol solution containing C8-TEOS/MTMOS ratios of A) 0.5 : 1, B) 1 : 1, C) 2 : 1. Reproduced with permission from [32].

MTMOS. These are shown in Fig. 6. The detection limits were comparable or slightly better than those obtained by using commercial SPME fibers. Mester et al. [33] have developed a SPME method to determine two methylated arsenic species, DMA and MMA, in human urine samples by GC-MS. Direct extraction of the methylarsenic compounds by SPME after thioglycol methylate derivatization was studied. Four different commercial SPME fibers were tested for the extraction of methylarsenic compounds, and the best results were obtained using the PDMS coating. Yates et al. [42] reported the preconcentration of neutral species by electrochemical control of SPME. Conductive polymer films were used as SPME elements for the direct and specific extraction of trace levels of AsB. Hydrophobic interactions between neutral arsenic species and an undoped polythiophene are responsible for the diffusion-controlled preconcentration. After absorption of analyte into the polymer matrix, the chemical properties of this conductive polymer were changed by applying an

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Guo et al. [44] combined SPME and electrochemically aided extraction for determination of trace level mercury. An SPME/EC fiber was made of a carbon steel wire with a 10-mm gold coating. Hg(II) ions were electrochemically extracted from aqueous solution, desorbed with a dedicated desorption system, and then detected by ion trap GC-MS. Hg(II) ions were detected in aqueous solution and mercury vapor in gas. Inorganic mercury and organomercury compounds were differentiated. Barshick et al. [45] developed a technique for the analysis of inorganic mercury salts in soils by GC-MS. Quantification of inorganic mercury was accomplished through a chemical alkylation reaction designed to convert an inorganic mercury salt into an organomercury compound prior to GC-MS analysis. Two alkylating reagents were investigated: methylpentacyanocobaltate(III) (K3[Co(CN)5 CH3]) and methylbis-(dimethylglyoximato)pyridinecobalt(III) (CH3Co(dmgH)2Py). Methylbis(dimethylglyoximato)pyridinecobalt(III) was found to be superior for this application. He et al. [46] analyzed organomercuric species in soils from orchards and wheat fields by CGC-AAS. After in-situ hydride generation, HS-SPME was carried out. Derivatization of polar organomercuric halides to their hydrides was undertaken in 0.1 M HAc-NaAc (pH 4) buffer by adding 1 mL of 6% KBH4; HS-SPME followed. Volatile derivatives were separated by GC and detected on-line by electrically heated quartz-furnace AAS. Mothes et al. [47] speciated organomercury compounds by SPME-GC-MIP-AED. The method was calibrated for aqueous and soil samples and the results indicated that www.jss-journal.com

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HS-SPME is a suitable tool for sampling and enrichment of Me2Hg and Et2Hg in those samples. GC hyphenated with MIP-AES detection was found to be a highly sensitive method for determining these species in the concentration ranges of ng/g and mg/L. A major problem was observed with the conservation of samples to avoid losses of the highly volatile Hg species during sampling and storage. Carro et al. [48] proposed a method for the speciation of mercury compounds by SPME-GC-MIP-AES in seawater samples. The Hg species were derivatized with Na tetraphenylborate, sorbed on a PDMS-coated fused-silica fiber, and desorbed in the injection port of the GC in splitless mode. Six variables, viz. sample volume, NaBPh4 volume, pH, sorption time, extraction-derivatization temperature, and rate of stirring were studied for optimization of the experimental conditions. Carpinteiro et al. [49] described a rapid and accurate method for simultaneous determination of MeHg+, Hg2+, monobutyltin (MBT), tributyltin (DBT), and tributyltin (TBT) in water samples by using a multicapillary GC column and MIP-AES detection. The samples are derivatized by ethylation and extracted by SPME. The species of both tin and mercury can be evaluated in the same chromato-


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graphic run by adjusting the auxiliary gas pressure and He cavity vent flow in MIP-AES. Other methods for the analysis of mercury in the presence of other metals such as lead, arsenic, and tin have also been reported [21, 24, 26, 29, 32, 37, 41].

6.4 Organotin Compounds Morcillo et al. [50] determined methyltin (MT) compounds in aqueous samples using SPME-CGC. In-situ derivatization was performed with sodium tetraethylborate and the analytes were adsorbed on a PDMS fiber in 2 – 35 min with stirring. Guidotti et al. [51] determined organotin compounds such as DBT, TBT, and tetrabutyltin (TeBT) in water samples by SPME and GC-MS. The compounds were derivatized with sodium tetraethylborate, extracted by a SPME fiber covered with a 100-mm PDMS coating, and desorbed into the splitless injector of a GC coupled with MS. It was used for the analyses of spiked river water. Lespes et al. [28] optimized SPME for the speciation of BT and PT in water samples. The analytical method consisted in in-situ ethylation followed by simultaneous SPME of the derivatives. The extracted sample was separated and detected by GC-FPD. Water samples were analyzed (Fig. 7) to verify the accuracy of the optimized

Figure 7. Chromatograms corresponding to the ethylationSPME-GC-FPD and to the ethylation-isooctane extraction-GCFPD analysis of a spiked synthetic water (100 ng/L of each compound): 1) MBT; 2) TPT; 3) MPT; 4) DBT; 5) TBT; 6) DPT; 7) TPT. Reproduced with permission from [28].

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method by comparing the results with those obtained using a conventional solvent extraction of the ethylated organotins. Wu et al. [52] identified organic tin compounds in contaminated lard by SPME-GC-MS. In the contaminated lard sample, six organic compounds were identified as raw materials or byproducts in the production of organic tin compounds. The method is simple and precise, and had not been previously reported in the literature. Vercauteren et al. [53] used HS-SPME-CGC-ICP-MS for determination of organotin in potatoes and mussels. NaBEt4 was used for derivatizing TPT and TCT to form sufficiently volatile compounds. Headspace extraction was performed for 10 – 20 min at 75 or 858C (depending on the type of sample) with a 100-mm PDMS fiber and the fiber was introduced into the ICP. Cardellicchio et al. [25] speciated butyltin compounds such as MBT, DBT, and TBT in marine sediments by using a mixture of HCl and methanol. The species were derivatized with NaBEt4 prior to HS-SPME extraction using PDMS. The derivatized organotin compounds were desorbed into the splitless injector and simultaneously analyzed by GC-MS. The method was optimized with respect to derivatization reaction and extraction conditions. Zhou et al. [54] investigated the occurrence of butyltin compounds in various environmental samples including water, sediment, marine products, and other commodities from China. The method involved hydrogenation coupled with SPME or Grignard derivatization, followed by GC-FPD analysis. The results demonstrated the ubiquitous presence of butyltin compounds. Aguerre et al. [55 – 58] used NaBEt4 as derivatizing agent in the monitoring of butyl- and phenyltin pollution in the environment by SPME-GC-FPD. SPME was applied for the first time to the simultaneous extraction of butyland phenyltins and the analytical performance was evaluated. This new method was employed for the analysis of various environmental samples such as natural aqueous samples, sediments, and sewage sludge [56]. A chemometric approach was adopted for organotin determination by CGC-ICP-AES. Typical chromatograms for ethylated BT and PT in isooctane are given in Fig. 8. Four factors were considered to assure success of the crucial part of the coupling which is passage of the analytes through the transfer line [57]. Hyphenation between solid-phase microextraction and CGC-ICPMS for the routine speciation of organotin compounds such as butyl- and phenyltin compounds in sediment and water samples was reported. All the factors were optimized by an experimental design approach to evaluate critical parameters influencing the overall analysis [55]. The performances of four specific detectors FPD, PFPD, MIP-AES, and ICP-MS for the speciation of organotins by using SPME-GC were

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Figure 8. Typical chromatogram obtained after injection of ethylated BT and PT in isooctane. Reproduced with permission from [56].

studied. ICP-MS is the most sensitive method. PFPD is also of significant interest [58]. Arambarri et al. [59] analyzed nine metals from ten superficial sediments from river estuaries of Gipuzkoa (North Spain). BT compounds were analyzed by AAS. The method involves the extraction of BT from sediments with a HCl-MeOH mixture followed by derivatization with NaBEt4. HS-SPME is preferred for the preconcentration using a PDMS fiber. The organotin species were analyzed by GC-FID. Lespes et al. [60] speciated organotin in French beans and potatoes cultivated on soils spiked with solutions or amended with sewage sludge. This was used to study its behavior in two vegetables cultivated on sandy soil spiked with solutions of either TPhT or TBT, and on a sludged soil. The method consists of acidic extraction of analytes from the plant material, followed by an aqueous ethylation-LLE-GC-PFPD. Le Gac et al. [27] reported an efficient and rapid method for the determination of organotin compounds by HSSPME followed by GC coupled to PFPD (Fig. 9). Six operating factors were considered to optimize the process (sorption on SPME fiber and thermal desorption in GC injection port). The accuracy was studied throughout the analysis of spiked environmental samples. Bravo et al. [61] identified sulfur interferences during organotin determination in harbor sediment samples by sodium tetraethyl borate ethylation and GC-PFPD. Chromatograms for LLE-GC-PFPD and SPME-GC-PFPD of differwww.jss-journal.com

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Figure 9. Typical chromatogram of spiked water obtained by aqueous ethylation-SPME-GC-PFPD. Reproduced with permission from [27].

Figure 10. Typical chromatogram obtained by LLE-GC-PFPD of an ethylated species from: a) a standard solution of OTC; b) an acidic extract of a harbor sediment sample (compound identification: 1) Hg2+; 2) MBT; 3) PPT; 4) BBT; 5) MPT; 6) TBT; 7) TeBT; 9) DOT, 10) TPT; 11) TOT, sulfur compounds); c) chromatographic separation of DBT and Et2S4 obtained with a new temperature program. Reproduced with permission from [61].

ent species from a standard solution of organotin compounds and harbor sediment sample are shown in Fig. 10 and Fig. 11. Because of the high toxicity of organotin compounds and the current regulations governing their applications, analytical methods usable in routine analysis are required. A speciation procedure based on NaBEt4 ethylation and GC-PFPD analysis proved to be suitable for organotin determination. Some matrix effects were observed due to the alkylation of elemental sulfur and coelution between the organotin compounds and some dialkyl sulfides. Chou et al. [62] used a procedure for the analysis of organotin which is based upon in-situ derivatization with Na tetraethylborate. This was followed by extraction by HS-SPME and separation and detection by GC-MS. The organotins were adsorbed on a

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PDMS-coated fused-silica fiber. The SPME experimental procedures involving the extraction of organotins in water were performed at pH 5, with extraction and derivatization proceeding simultaneously at 458C for 30 min in a 2% Na tetraethylborate solution. Desorption was carried out in the splitless injection port of the GC at 2608C for 2 min. The proposed method was tested by analyzing surface seawater from harbors on the Taiwanese coast for organotin residues. Girona et al. [63] analyzed BT, PT, and HT by SPME-CGC after derivatization with sodium tetraethylborate. The analytes were extracted using a PDMS fiber. The fiber was immersed for 30 min in the sample solution with constant stirring to assure proper extraction of the analytes. www.jss-journal.com

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Figure 11. SPME-GC-PFPD chromatogram obtained from acidic extracts of a harbor sediment sample. Reproduced with permission from [61].

It is reported that organotins can also be analyzed in the presence of other metals [15, 21, 24, 26, 29, 32, 35, 37, 38, 49].

6.5 Organoselenium compounds Landaluze et al. [64] monitored organoselenium compounds in the production and gastric digestion of selenized yeast. Determination of volatile species of selenium was achieved by coupling SPME for preconcentration and sample-matrix separation with CGC-MIP-AES. The MC column was operated at low temperatures. The method was optimized, using a chemometric approach, with respect to the detection of organoselenium species such as dimethyl selenide (DMS), dimethyl selenide (DES), and dimethyl diethyl selenide (DMDS). HS-SPME was used for sampling. The results of the yeast enrichment process demonstrate transformation of inorganic selenium into volatile organic species. Dietz et al. [65] applied SPME-MC-GC coupled to different detection systems to volatile organoselenium speciation in yeast. The methods compared for detection were ICPMS, MIP-AES, and AFS. All the detectors were suitable, with the highest sensitivity being obtained for MIP-AES detection, with detection limits of 0.57, 0.47, and 0.19 ng/mL for DMS, DES, and DMDS, respectively.

7 Conclusions Solid phase microextraction (SPME), developed in 1990, is now an established sampling technique for the extraction and on-line desorption of metals and organometallic species. The sensitivity and selectivity are dependent on the type of fiber and on the detector used with the chromatographic technique. The more advanced technique of in-tube SPME, based on tubular fused-silica capillary columns, is also attracting the attention of many workers. Thus it can be concluded that SPME offers great advantages over classical sampling techniques, which are time consuming and require large amounts of samples and solvents. Thus, the future of SPME appears highly promising.

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