Chapter 2 Sample preparation techniques for elemental speciation studies

Chapter2

Sample preparation techniques for elemental speciation studies Joanna Szpunar, Brice Bouyssiere and Ryszard Lobinski

2.1 INTRODUCTION The growing interest in trace element speciation over the last two decades has been reflected in the development of a number of hyphenated (coupled, hybrid) analytical techniques which are able to discriminate among the different forms of an element [1-6]. These techniques usually show an outstanding performance for standard solutions but often fail for a real sample. It is now obvious that the key to a successful speciation analysis lies in the revival of old-fashioned wet-chemical sample preparation methods, which were almost lost with the advent of powerful quasi-direct multielement analytical techniques such as ICP-AES and ICP-MS [7]. In speciation analysis, wet sample preparation methods have a unique role to play because of the low analyte concentrations involved (often below 0.1 pg/g or 0.1 ]jg/l), the fragility of the analytes and the need for preservation of the organometallic moiety throughout the procedure, and the strict requirements posed by the coupled system in terms of the sample volume, polarity and matrix acceptable by the instrumental setup. Another distinct trend is automation, which implies the development of faster, possibly single-step, and efficient sample preparation procedures. The procedure for sample preparation depends on the analytical technique to be used and on the sample type to be analyzed. The polarity or volatility of the analyte determines the chromatographic technique to be chosen for the separation of the species prior to detection. The separation technique, on its turn, sets the requirements for the analyte solution resulting from the sample preparation procedure. Comprehensive Analytical Chemistry, Vol. XXXIII J.A. Caruso, K.L. Sutton and K.L. Ackley (Eds.) © 2000 Elsevier Science B.V. All rights reserved

7

Plant and animal biochemistry, ecotoxicology, nutrition Organometalloid

Phytochelatins

~~speesn . .. arsenobetaine,

Macrocycles -. chlorophyll derivatives, cobalamiiines arides ...... Pb Sr Ba, Ca, Mg . Metalloenzymes Zn, Mo, Co

Cd, .... Cu, Zn, Co, As

arenocholine, selenOaminoacids Metallothioneins Cd, Cu, Zn Redox states: As(lll)/As(V),Cr(lll)/Cr(VI) Se(lV)/Se(VI), Sb(ll)/Sb(V) Fe(ll)/Fe(lIl)

/

Metal

and

metalloid species

Alkylmetals: 3

Metallodrugs Pt, Ru, Ti Nucleic acids Cr, Ni, Pt, Ru

+

I

5

3

MeSn( -n), BuSn(3) ,.... Ph,Sn(.n), MenGe4 ...

Me 2Hg, Et2Hg, MeHg MenEtmPb(4-rl-)+ MeCd+ Me2Cd

5.

a

+

Metalloporphyrins Ni, V, Fe, Ga

Catalytic mixtures Ni, Rh, Ru

Transport proteins .. Al, Cu. Zn, Fe

I

Organomercury and -arsenic Shale oil, petrol, natural gas condensates ",..

Industrial chemistry Fig. 2.1. Species and fields of interest in speciation analysis.

Gas chromatographic techniques would require a limited volume of a non-polar solvent free of suspended particles containing volatile thermally stable species. An aqueous solution of thermodynamically stable and kinetically inert species (organometallic or metal complexes) free of suspended particles will be required for liquid chromatographybased techniques. In terms of analytes, three major areas can be distinguished: speciation of redox forms, speciation of organometallic [containing a carbon-metal(metalloid) bond] and speciation of metal complexes. Figure 2.1 summarizes the principal species for speciation analysis. Gas chromatography is the preferred separation technique for alkylelement species whereas liquid chromatography is predominantly used in all the other cases. The choice of sample preparation procedure is also determined by the matrix and the preconcentration factor that needs to be achieved to eliminate the discrepancy between the concentration of the analyte in the sample and the detection limits of the analytical setup. A sample preparation procedure for speciation analysis usually requires a number of steps (Table 2.1) including filtration, preconcentration of 8

TABLE 2.1 Glossary of the most common steps in sample preparation for speciation analyses Filtration

Solubilization

Leaching

Preconcentration

Clean-up

Derivatization

is used to separate particles in gases and aqueous samples. Usually a 0.45 pm filter is used but 0.2 pm filter is necessary prior to HPLC. is applied to bring biological materials into solution. It can be achieved by alkaline, acid or enzymatic hydrolysis. Following the solubilization, an aqueous solution of species is obtained but the matrix is not eliminated. or solid-liquid extraction is applied to extract analytes species from solid samples (soil, sediment or biological tissues). Leaching is the most popular for speciation of metal complexes since pH neutral leaching agents. Leaching is also the preferred methods for the analyses of soil and sediment samples that cannot be solubilized without the destruction of analytes. is necessary to increase the concentration of analysed species in the solution introduced on a chromatographic column in comparison with that present in an analysed sample. The techniques used include cryofocussing, gas-solid extraction (trapping) (for analytes already in the gas-phase), solid-phase microextraction (SPME) for analytes in gas or liquid phase, liquid-solid extraction (sorption) for analytes in water. Preconcentration can be achieved by solvent extraction and evaporation of the leachate or extract. is the removal of the matrix components (fats, proteins, highboiling point hydrocarbons) that, if co-introduced on a chromatographic column would lead to their destruction or degradation of separation properties. Cleanup is usually realized by low-resolution chromatographic separation with a mechanism different to that employed for the analytical separation (e.g. passing through a Cs column of polar compounds to be separated by anion-exchange) is the process of the controlled conversion of species originally present in a sample into forms with improved chromatographic yield or separation coefficient. The most popular is derivatization of ionic or highly polar species into non-polar that can be readily separated by GC (e.g. Grignard derivatization).

analytes, solubilization or leaching in the case of solid samples, cleanup prior to chromatography, and sometimes derivatization of analyte species to improve their chromatographic behavior. It is essential that all these steps be carried out with a maximum (quantitative) efficiency and that the original species are not degraded. The number of steps necessary and their duration affect the duration and tediousness of an 9

analytical procedure and should be kept minimal. Only some energyrelated samples (shale oil, petroleum, gasoline) containing low polar analytes can sometimes be analyzed directly, usually after dilution. This chapter discusses sample preparation methods in speciation analysis starting from the sample collection and ending with the injection of a solution of analyte species on a chromatographic (gas or liquid) column. Emphasis is put on procedures that require a minimum of time and a small number of operations, keeping in mind the automation of the sample preparation process.

2.2 SAMPLE COLLECTION, PRETREATMENT AND STORAGE Many errors arise at early stages of the analytical procedure (sample collection and storage), but little is known about the effect of particular factors on speciation analysis. Precautions to avoid contamination need to be taken during sampling, especially in the remote environment, in particular for organolead and mercury analyses. Initial cleaning of containers is essential, especially if the same containers are going to be used for samples with different concentrations of the analytes. Generally, sampling and storage protocols, recommended for the analysis of a particular type of sample for trace metals or volatile organic contaminants, should also apply to organometals. Some specific requirements are discussed below. 2.2.1 Atmospheric and gas samples Samples include atmospheric or workplace air, gases emitted from domestic waste deposits and from laboratory experiments (e.g. studies of biomethylation by sediments) [8-11]. Species of interest usually include tetraalkyllead compounds (MenEt4 _Pb), methylselenium compounds (e.g. Me 2Se, Me 2Se2), and some organomercury compounds (MeHg + , Me2 Hg). The compounds may be present in the gaseous phase or be associated with the particulate (aerosol) phase. The distinction between gaseous and particulate species in air is purely arbitrary, and depends on the type of filter used. Analytical procedures for sampling air are usually integrated with an enrichment step carried out in the field. Contamination is reduced by capping the sampling column immediately after sampling and by excluding air from the column. Older approaches included a series of 10

columns each containing a species-selective sorbent, e.g. a glass wool filter for the removal of particulate Hg forms, SE-30 coated chromosorb for the removal of HgC12, NaOH-pretreated Chromosorb for the removal of MeHg*, Ag-coated beads for the removal of Hg °, and Au-coated beads for the removal of Me 2 Hg [12]. Proving that the specificity of adsorbents is valid for all natural conditions is hard. A better approach is trapping the maximum number of species on one sorbent followed by their chromatographic analysis with element-selective detection. The typical sampling train consists of a 0.45 um filter which separates the gaseous phase (being subject to analysis) from the particulate matter (aerosol), followed by an empty cold trap (-40°C) to remove the water vapor, and the actual sorbent trap (often cooled with liquid nitrogen) for the analyte compounds [5,8]. The whole gas train should be deactivated on a regular basis with a silanizing agent to minimize adsorption on active sites. The sorbent or cryogenic trap is brought to a laboratory for the analysis. Samples can apparently be preserved in liquid nitrogen for several days [8]. The analytes retained on the sorbent can be desorbed with a moderately polar solvent, such as ethylacetate or toluene, or released by thermal desorption onto a GC column. Often the trap itself acts as a chromatographic column [8]. Much work is still required to assure the reliability for sampling metal species. Caution is recommended in interpreting results obtained under atmospheric conditions because of the possibility of degradation by cotrapped atmospheric components, e.g. ozone. An FeSO 4 filter is sometimes included to remove atmospheric ozone that, when retained together with the analyte compounds, could promote the degradation of the latter. Decomposition during sampling may be compensated for by the use of an internal standard of deuterated Me 4 Pb and Et 4 Pb [13] or by compounds marked with stable metal isotopes. Difficulties with the proper addition of this standard to the sample prior to trapping occurred. Atmospheric particulate matter collected on a glass fiber filter are dealt with as described below for sediments. 2.2.2 Water samples The initial cleaning of the containers is important to avoid contamination, especially if the same items are used for samples with different analyte concentrations. An efficient way of cleaning consists of a simple soaking of the glassware in hot concentrated HNO0 to decompose 11

organometallic compounds previously adsorbed. The sampling protocols extensively described in the literature on ultratrace analysis can usually be applied. Acidification of samples is recommended to prevent losses by adsorption on the container walls. The need for filtration prior to analysis is dependent on the level of particulate suspended matter. In unfiltered samples, changes in the distribution of organometallic compounds may occur with time due to adsorption and desorption processes at the particles' surface. Filtering ensures homogeneity of the analyzed sample but may lead to losses of volatile, e.g. Me 2Se species. In general, filtration is recommended for samples with high particulate load such as urban deposits, river, estuarine and shallow coastal waters, and eutrophic lakes; it can be avoided for sea water, tap water and atmospheric deposits from remote environments. The preservation of analytes in their authentic state prior to analysis may present difficulties. The less polar the organometallic species, the more likely is its sorption onto glass walls leading to the decomposition of the compound. Ionic organometallic species are not adsorbed to an appreciable extent. Organometallic compounds are known to decompose in solution in a light-induced process promoted by microorganisms, suspended solids and various impurities [14]. Tetraalkyllead species were found to be the least stable. No noticeable change of ionic alkyllead species in water samples stored in glass bottles at 4°C in the dark was observed for a period of up to 1-3 months [14]. Decomposition rates depend dramatically on the origin and composition of the sample, and very little is known about the stability of dilute solutions (below 10 ng 1-1). The most appropriate way of storage is to deep-freeze samples immediately on collection to minimize any bacterial or enzyme degradation, loss through volatility, or contamination of the sample. Water samples frozen in polycarbonate containers for tributyltin were shown to be stable for 2-3 months [15]. The container material was critical in minimizing surface adsorption [16]; PTFE provoked substantial losses. Immediate extraction of analytes into an organic solvent can minimize losses or degradation originating from the sample container [17]. For dissolved gaseous species, such as e.g. Hg ° , Me 2 Hg or methylselenium storage generally leads to losses and/or contamination. Samples containing these analytes should be purged immediately onto a trap, which can provide stable storage. The storage of MeHg in aqueous solutions is subject to disagreement. MeHg-spiked tap water 12

samples stored in Teflon bottles were stable for at least one month after quick freezing in liquid N 2 followed by storage at -80°C, and finally storage at 0-4°C in the dark. Samples stored at room temperature after acidification with 0.1% HC1 showed a rapid decrease (80% after 2 days) of the MeHg concentration. The effect of storing seawater samples for mercury speciation was studied [18]. The stability of MeHg in deionized water was found to depend on its concentration, the container material and the storage temperature, whereas light seemed to have no effect [19]. MeHg solutions stored in Teflon containers at pH 6 were stable for at least 20 days; in glass containers good results were obtained after acidification (1% HNO3 ) and storage at 0-5 0 C. The storage characteristics of selenate, selenite and selenomethionine in low and high ionic strength water [20], iodine species [21], chromium immobilized on alumina microcolumns [22], arsenic species in fresh and sea water [23] have been investigated. An overview on the stability of chemical species during storage, based on results from projects carried out by the Commission of the European Communities Measurements and Testing Programme (formerly BCR), aimed at improving the quality of speciation analysis, is given in Ref. [24]. 2.2.3 Sediments and soils Particulate matter suspended in natural waters can be collected by filtration using a 0.45 m filter. For representative samples, however, very large volumes should be filtered, which brings a risk of filter clogging. Therefore, continuous-flow centrifugation is recommended. Relatively little work has been devoted to the behaviour of organometals in suspended matter and sediments during storage. Different methods of preserving soils and sediments, i.e. wet at room temperature, wet at lowered temperature, frozen, freeze-dried, oven-dried, microwave oven-dried and air-dried at room temperature were considered [25]. No method preserved the metal species. Air drying was the most convenient method as it minimized the storage problem and was readily homogenized and subsampled. The recommended soil drying procedure was as follows: soil (1-2 kg) was spread thinly on a polyethylene sheet on aluminium racks in a fan-ventilated room and heated to 25°C for a few days [25]. Larger aggregates were broken down by gentle rolling on a thick polyethylene sheet with a wooden roller and the soil was sieved through holes 2 mm in diameter. The <2 mm diameter soil particles were stored in screw-capped glass bottles until needed. 13

Sampling and storage of sediment samples for organotin speciation has been discussed [26]. Freezing and lyophilization were suitable to preserve butyltin for 12 months whereas worse stability was observed for phenyltin. Regarding methylmercury, the main source of instability is due to bacteria, either by demethylation [27] or formation of volatile dimethylmercury [28]. It was found that methylmercury was not affected by y-irradiation and that this procedure could be used for the stabilization of sediment samples [29]. Avoiding contamination and loss is followed by sections on soils (field sampling, and sub-sampling of air-dried soil) and sediments (location of sampling points, kinds of sample and sampling apparatus for suspended sediments, bottom sediments and interstitial water, measurement in situ, and pre-treatment and storage of the sample) was discussed [301. An extensive discussion on the stability of candidate CRM sediment samples in terms of concentrations of organometallic compounds can be found elsewhere [6]. 2.2.4 Biological materials Protocols recommended for the determination of element traces in biological materials also apply to speciation analysis. However, the need for rapid analysis or freezing the samples needs to be emphasized [31], otherwise the enzymatic activity and natural proteolysis and autolysis processes will continue after sampling and can alter the speciation. The lipophilic character of non-polar compounds favors their accumulation in particular tissues of living organisms so the dissection of the parts of interest and homogenization must precede the analysis. Sample tissue or plant material can be pulverised in liquid nitrogen to break up the matrix. Blood samples were hemolyzed by freezing (-20°C) for at least 24 h [32]. Storage for extended periods under such conditions was found not to affect the stability of organolead initially present. Storage of samples in daylight should be avoided due to potential losses especially of ethyllead species [33]. A high rate of degradation of tri- and dibutyltin in mussel tissue was observed at 40°C, instability was also noted at 20°C and at 4°C; -20°C was recommended for long-term storage; phenyltin compounds were found to be unstable [6]. No instability could be demonstrated for total mercury and methylmercury content in fish tissue stored for 12 months at -20°C up to 40°C [6]. Freeze-drying caused a loss of methyltin [34]. The stability of organometallic species in candidate reference biomaterials is extensively discussed elsewhere [6]. 14

2.3 SAMPLE HANDLING PRIOR TO GC CHROMATOGRAPHIC

ANALYSIS

Gas chromatography requires an analyte species be presented as a nonpolar, thermally stable compound in a non-polar solvent or in a narrow (cryofocussed) band. The sample preparation procedure should therefore include either a step transferring analytes from the aqueous solution (water sample, leachate or solubilizate) into an organic solvent (solvent extraction) or a step of formation of a narrow analyte band on a chromatographic sorbent by purge-and-trap or solid-phase microextraction. The derivatization step to yield a thermally stable species may either follow the solvent extraction step (Grignard derivatization) or precede it (derivatization with NaBH4 or NaBEt4 ). An additional preconcentration step by sorption or evaporation is necessary where ultratrace amounts of analytes are determined and/or poorly sensitive detection techniques are used. A schematic layout of sample preparation methods for speciation analysis of organometallic species in aqueous samples by capillary GC with element selective detection is shown in Fig. 2.2 [35]. Sample

Analyt

Complexing

Separation

I

Deivatization

Preconntation

| Detection

Fig. 2.2. Sample preparation techniques for speciation analysis by gas chromatography with element selective detection. 15

preparation methods fall into two basic categories: methods based on in-situ derivatization of the analytes followed by their purging and cryotrapping, and methods based on the extraction (solid phase or liquid-liquid) of the analyte species, either native, or as non-polar complexes (with DDTC, dithizone or tropolone), or derivatized beforehand in the aqueous phase. 2.3.1 Aqueous samples The analysis is preceded by the separation of analytes from water, which is usually combined with their preconcentration, and often with their derivatization. Several possibilities exist as described below. 2.3.1.1 Sorption (solid-phase extraction) Solid-phase extraction (SPE), which is becoming increasingly popular for sample preparation in organic analysis, found its way to speciation analysis for organometals [36-41]. The analytes are extracted by sorption, eluted with a small amount of an organic solvent and derivatized. The advantages of SPE over liquid-liquid extraction include a higher enrichment factor, lower solvent consumption and risk of contamination, and the ease of application to field sampling and automation. Only filtered samples can be analyzed, which may constitute a considerable drawback. SPE cartridges in a variety of configurations and sizes have been used but microcolumns proved to be the best choice with respect to low dead volumes and low consumption of the solvent [36]. C18 extraction disks, reported to be well suited for the trace enrichment of organics in environmental waters, did not show satisfactory retention of native di- and monobutyl(-phenyl) tin species [36]. Sorbents with immobilized chelating reagent (e.g. dithiocarbamate) have widely been used [37-40]. A sampling and storage technique for speciation analysis of lead and mercury in seawater based on the sorption on a column packed with a dithiocarbamate resin was proposed [37]. An interesting curiosity is the use of biomaterials, e.g. dried yeast (Saccharomyces cerevisiae) cells for enrichment of methylmercury from river water [41]. 2.3.1.2 Solid phase micro-extraction Solid-phase micro-extraction is a preconcentration technique based on the sorption of analytes present in a liquid phase or, more often in a headspace gaseous phase, on a microfiber coated with a chromato16

graphic sorbent [42,43]. When the equilibrium is reached, the fiber is transferred to a GC injector by means of a microsyringe, and the analytes are thermally desorbed inside the heated injector onto a capillary chromatographic column. A method based on SPME on a fiber coated (100 pm) with poly(dimethylsiloxane) was developed for the determination of tetraethyllead in water [44]. A water sample was also subjected to headspace SPME of ethylated organometallic species onto polydimethylsiloxanecoated fused-silica fibres for sensitive simultaneous determination of organomercury, -lead and -tin compounds [45]. For aqueous phase solid-phase micro-extraction, methylmercury in river water samples was in situ derivatized using NaBEt4 and a silica fiber coated with poly(dimethylsiloxane) was placed into the solution and when equilibrium was reached, the fiber was thermally desorbed [46]. Methyltin chlorides and tetramethyltin were adsorbed from acetone solution onto methylsilicone coated fibers and transferred into a headspace vial for equilibration with NaBH4 solution [47]. 2.3.1.3 Solvent extraction Solvent (liquid-liquid) extraction can be directly applied to non-filtered samples with complex matrices and allows the direct transfer of analytes into a non-polar organic solvent that can be analysed by GC. Non-polar species, e.g. tetraalkyllead, are quantitatively extracted from water saturated with NaCl into a 20 times smaller amount of hexane [48]. The same applies to some "ionic species" with the marked covalent character (e.g. triphenyltin, tributyltin, triethyllead, methylmercury) but the use of water-immiscible solvents with polar character (toluene, ethylacetate) is advised. The properties of organometallic species with a smaller number of organic substituents (e.g. monobutyltin) do not allow their quantitative extraction by any organic solvent. In order to transfer such compounds into an organic phase, the formation of extractable chelate complexes or non-polar covalent compounds in the aqueous solution prior to extraction is necessary. Diethyldithiocarbamate (DDTC) is the most often used reagent to form the chelate complexes with organometallic species. The extraction is followed by a derivatization step, usually by a Grignard reaction. Extraction of the complexes of ionic organolead species with DDTC from pH 6-9 into hexane [49-52], organotin [53-56] and organomercury [57,58] was found to give quantitative recovery. Dithiocarbamates are not as light sensitive as dithizonates making the handling easier 17

and the procedure more reliable. The high selectivity of the hexanetetramethyldithiocarbamate extraction system for ionic alkylleads over Pb 2+ facilitates greatly the determination of these analytes in matrices contaminated with high levels of Pb2+ [59]. Inorganic interferents can be efficiently masked with EDTA [49-52]. For organotin, tropolone has extensively been used [56,60-63] in addition to DDTC. An alternative to chelate extraction is the formation of extractable non-polar species in the aqueous phase by hydride generation [64,65], alkylation with NaBEt 4 [36,66-70] or NH4 BBu4 [71] or arylation with NaBPh 4 [66,72]. The resulting species should have a boiling point exceeding by 20°C that of the solvent to be used for their extraction in order to enable the subsequent gas chromatographic separation. Also, it is important that the nucleophilic substituent be different from organic substituents of the species to be analyzed. For example, ethylation will fail for Et2 Pb 2+ and Et 3Pb+ since it would lead to the identical species (Et 4 Pb) which is also the product of Pb2+ ethylation. Preconcentration factors in solvent extraction are generally low (typically 1:50 up to 1:250). The methods are rapid, work well for less volatile analytes and are relatively robust in terms of interferences. 2.3.1.4 Steam distillation Steam distillation was evaluated as a technique for the separation of methylmercury from natural water samples [73-76]. This technique was found to give higher and more reproducible recoveries than other extraction techniques but under some conditions it could be responsible for artificial methylation of Hg2+ [74,75]. The technique is rather slow; the addition of ammonium pyrrolidine dithiocarbamate (APDC) was found to improve recovery (to 85% for seawater) and to eliminate the codistillation of inorganic mercury. The method was found to be artifact-free and to be comparable to nitrogen-assisted distillation with the added advantage of increased samples throughput [76]. 2.3.1.5 Liquid-gas extraction (purge and trap) This technique consists of bubbling an aqueous solution with an inert gas (nitrogen or helium) to extract the non-polar volatile species into the gas phase. Some species can be extracted directly (e.g. tetraalkyllead, tetramethyltin, dimethylmercury) but others need to be converted in volatile hydrides or ethyl derivatives. Purge-and-trap methods usually follow hydride generation, which is the oldest but still the most popular method for volatilization of As, Ge, Sn, Sb and of their 18

methyl species [77-80]. Purging of less volatile tributyl- or triphenyltin hydrides is negatively affected by condensation problems, which may lead to non-quantitative recoveries of the analytes. In general, hydride generation is prone to matrix interferences and hydrides are relatively reactive, tending to decompose when subject to harsh instrumental conditions. A rapidly developing alternative for the production of volatile derivatives is ethylation using NaBEt4 . Ethylation was developed for ionic mercury [81,82], lead [82,83], tin [82] and selenium [84] species and enjoys a continuously increasing number of applications. Ethyl derivatives of butyl- and phenyltins are not sufficiently volatile to be efficiently purged. Furthermore, this technique is not selective for ethylmetal species. The purged species are subject to moisture removal (as in the case of the analyses of gases), are cryotrapped, and released onto a packed or capillary column. This method offers high preconcentration factors and allows the introduction of fairly clean samples onto a GC column. Interferences, especially with hydride generation, and the risk of condensation with less volatile analytes are the major shortcomings. 2.3.2 Solid samples Solid samples of interest for speciation analysis can be divided in two major categories: sediment and soil samples and biological materials. Organometallic compounds are apparently not involved in mineralogical processes in sediment and soil and bind onto the surface. Therefore, the complete dissolution of the sample prior to analysis is not considered necessary. On the contrary, organometallic compounds may be incorporated in tissues of a living organism. Hence, solubilization of a biological material prior to separation of the analytes is mandatory even if in particular cases some success can be obtained by leaching. Once the analyte species are brought into solution, the latter is treated similarly to water samples as described above. 2.3.2.1 Leaching methods The basic approach to release organometallic compounds from the sediment involves acid leaching (HCl, HBr, acetic acid) in an aqueous or methanolic medium by sonication, stirring, shaking or Soxhlet extraction with an organic solvent [53,55,85-94]. In order to increase the extraction yield, the addition of a complexing agent (tropolone, DDTC) is preferred. 19

A modification of the leaching procedures is distillation of methylmercury. A sediment, soil or biological tissue sample is suspended in an acidic solution, and the mixture is distilled at elevated temperatures (ca. 180°C) with nitrogen [95-99]. The distillate is collected in an icecold container. The complexation reagent, sodium pyrrolidinedithiocarbamate, may be added to improve the extraction recovery. Artifact formation of methylmercury during procedures for its extraction from environmental samples was studied [100]. For soil and sediments, the need for a cumbersome sample preparation step is the basic weakness of the whole analytical procedure. Indeed, the majority of procedures reported have not only been extremely time-consuming, taking from 1 h to 2 days, but also were usually inefficient in terms of analyte recovery and, in general, unreliable. As shown by Chau and co-workers [101], only three out often sample preparation methods described in the literature for the analysis of sediments were able to recover more than 90% of Bu 3 Sn +, whereas none of them was able to recover monobutyltin (BuSn 3+) in a reproducible manner. In general, the more polar the species to be extracted, then there will be less recovery, a longer leaching procedure, and a higher demand for accelerated extraction techniques (supercritical fluid extraction, accelerated solvent extraction or microwave assisted leaching). 2.3.2.2 Solubilization of biological samples prior to speciation analysis A suitable digestion (hydrolysis) procedure should allow for the complete destruction of the matrix while the organometallic moiety remains unchanged. Three principal approaches have been followed: (1) acid hydrolysis with HCl [102]; (2) alkaline hydrolysis with methanolic NaOH [103,104] or with tetramethylammonium hydroxide (TMAH) [105,106]; and (3) enzymatic hydrolysis [107-109]. 2.3.2.3 Enhanced techniques: supercriticalfluid extraction and accelerated solvent extraction Substantial progress towards faster and potentially automated speciation analysis of sediments is offered by supercritical fluid extraction [110-119]. Equipment cost is, however, high, the extraction step still requires 10-50 minutes and the recoveries of many species are far from being quantitative. A recent alternative is accelerated solvent extraction (ASE) [120,121]. ASE is based on performing static extractions at 20

elevated temperatures and pressures. The pressure can be programmed without the use of elevated solvent temperatures that could lead to decomposition of thermally unstable compounds. 2.3.2.4 Microwave-assistedprocesses Microwaves are high frequency (2.45 GHz) electromagnetic waves which are strongly absorbed by polar molecules (e.g. water or mineral acids) and which interact only weakly with non-polar solvents. Absorption results in dielectric heating; the heat appears in the core of the target sample. The well known advantages of microwave heating such as absence of inertia, rapidity of heating, efficiency and ease of automation have made it widely used for accelerated extraction of polar compounds into a non-polar or weakly polar solvent. The preservation of the organometallic moiety is the prerequisite of a successful leaching/digestion procedure prior to speciation analysis, and this can be achieved by a careful optimization of the microwave conditions [1221. In contrast to common high temperature and pressure acid procedures, a focused low power microwave field is preferred for extraction of organometallic species from the matrix. The carbonmetal bonds remain intact. A schematic of a focused microwave system, which has been used in the majority of speciation studies is shown in Fig. 2.3. Microwave-assisted leaching was shown not only to reduce the time necessary for leaching of organometallic compounds from soil and sediment samples, but also to increase the recovery of the most difficult

or

magnetror

on

focused microwave field

Fig. 2.3. Schematic of the focussed microwave digestor. Model 301, PROLABO, France.

21

to extract compounds. The time necessary for the quantitative extraction of organotin [123-126], organolead [127,128] and methylmercury [129,130] was reduced to 2.5-3 min. The values obtained for the extraction recoveries of monobutyltin from CRMs and candidate CRMs by these techniques are among the highest reported in the literature. Microwave fields can also accelerate leaching of analyte compounds from biomaterials [131,132]. When a suitable reagent is used (e.g. tetramethylammonium hydroxide) the biological tissue can be solubilized within 2-3 min instead of 1-4 h [124,125,133-135]. An even more attractive alternative is the integration of the solubilization, derivatization and extraction steps into a one-step procedure. Hydrolysis with acetic acid carried out in a low-power focused microwave field in the presence of NaBEt4 and nonane was shown to shorten the sample preparation time for the CGC-MIP AED determination of organotin compounds in biological materials to 3 min [136]. 2.3.3 Organic samples (gas condensates, shale oils, gasoline) This category includes various samples, of which the most often analyzed are: (1) fuels, for alkyllead [137,138] and manganese carbonyl compounds [139,140]; (2) crude oils, for metalloporphyrins [141-143], and natural gas condensates for mercury species [144-146]. The analytes are volatile and thermally stable so that samples can usually be directly injected onto a GC column. Gasoline is often diluted ten times to avoid the interference with hydrocarbons and the resulting plasma saturation. The detection limits for metals in oils and gas condensates are high in comparison with those for other samples. This is due to the hydrocarbon matrix which gives rise to background interference when it enters into the plasma at the same time as a species of interest. Moreover, carbon compounds can overload the plasma discharge, which has limited thermal energy, and hence reduce the excitation ability. The determination of metalloporphyrins in crude oils is further complicated by their low concentration levels, wide range of molecular weights, the complicated isomerism and the complexity of the crude oil matrix [141-143]. Hence, a sample pretreatment step is often necessary. In order to counteract the degradation in GC performance due to the accumulation of nonvolatile residues from the crude oil, removal of the asphaltene and pigment materials has been recommended. This step also enables reduction of the maximum elution temperature for a 22

sample, thus protecting the column and reducing the bleed and stabilizing the detector [143]. Other benefits of a sample pretreatment include group isolation of metalloporphyrins and non-porphyrins for GC and preconcentration of metalloporphyrins for trace analysis [143]. An elegant solution to avoid the interference from hydrocarbons was proposed for speciation analysis of mercury in gas condensates [144]. Mercury containing peaks are collected from the column eluate by an amalgamation trap. They are subsequently released into the plasma as Hg° in a flow of helium. Methylmercury and labile ionic mercury can be extracted as complexes with cysteine into the aqueous phase to eliminate the matrix and then re-extracted into hexane prior to gas chromatography [145]. A direct approach to speciation analysis of mercury in gas condensates was recently proposed [146]. 2.3.4 Derivatization techniques in speciation analysis A number of native organometallic compounds which are volatile enough to be separated by GC exist including tetraalkyllead species (MenEt 4 _nPb), methylselenium compounds (e.g. Me2 Se, Me 2Se2), some organomercury compounds (MeHg', Me 2Hg) as well as naturally occurring metalloporphyrins. As indicated above, these compounds can either be readily purged with an inert gas or extracted into a non-polar solvent and, subsequently, chromatographed by thermal desorption, packed column or capillary GC. The majority of organometallic species exist in quasi-ionic polar forms which have relatively high boiling points and often poor thermal stability. To be amenable to GC separation, these species must be converted to non-polar, volatile and thermally stable species. The derivative chosen needs to retain the structure of the element-carbon bonds to ensure that the identity of the original moiety remains conserved. The most common derivatization methods include: (1) conversion of inorganic and small organometallic ions into volatile covalent compounds (hydrides, fully ethylated species) in aqueous media; (2) conversion of larger alkylmetal cations: e.g. RnPb (4 ')n with Grignard reagents to saturated non-polar species; and (3) conversion of ionic species to volatile chelates (e.g. dithiocarbamate, trifluoroacetone) or other compounds. The three methods are fairly versatile in terms of organometallic species to be derivatized and the choice depends on the concentration of interest, the matrix and the sample throughput required. 23

Frequently, the derivatives are concentrated by cryotrapping or extraction into an organic solvent prior to injection onto a GC column. 2.3.4.1 Derivatizationby hydride generation Several elements (Hg, Ge, Sn, Pb, Se, Te, Sb, As, Bi, and Cd) can be transformed into volatile hydrides, forming the basis of their determination [77-80]. The usefulness of this procedure for speciation analysis, however, is severely restricted by either the thermodynamic inability of some species to form hydrides, or by considerable kinetic limitations to hydride formation. Nevertheless, the technique is still essential for some classes of compounds. Selenium, As, and Sb readily form hydrides only in their lower oxidation states; the higher states need to be reduced beforehand. Thus, all inorganic species of these elements form eventually the same hydride (SeH 2, AsH 3, and SbH3 , respectively) precluding simultaneous chromatographic speciation. Methylarsonic and dimethylarsinic (or stibonic) acids can be discriminated in one GC run upon hydride generation producing volatile MeAsH 2 and Me2 AsH (or MeSbH2 and Me 2SbH, respectively). Trialkyllead species form stable hydrides whereas dialkylleads are non-reactive. Mercury(II) and methylmercury [78], as well as germanium and methylgermanium species [80] can be converted to gas chromatographable hydrides. Hydride generation has enjoyed the largest interest for organotin speciation analysis because of its capability for the simultaneous determination of ionic methyl and butyl species in one chromatographic run [79,88,109]. Hydride generation with NaBH 4 is prone to interferences with transition metals which affect the reaction rate and analytical precision [77]. 2.3.4.2 Derivatization with tetraalkyl(aryl)borates The vulnerability of hydride generation to interferences in real samples and the restricted versatility can, to a certain degree, be overcome by replacing NaBH4 by alkylborates. The most common derivatization procedures rely on ethylation with sodium tetraethylborate (NaBEt4 ) which is water soluble and relatively stable in aqueous media. The use of NaBEt4 in speciation analysis has been reviewed [70]. Methyl-, butyl- and phenyltin compounds react readily with NaBEt4 to form thermally stable gas-chromatographable species. Whereas methyltins can be purged upon derivatization, other species need to be extracted because of their poorer volatility. All alkyllead species also readily react, but only methyllead species can be unambiguously dis24

criminated as the derivatization of ethyl- and inorganic lead will lead to the formation of the same product: PbEt4 . Methylmercury and inorganic mercury can be determined in one run upon purge-and-trap preconcentration. Derivatization of various selenium species was demonstrated but showed poor potential for simultaneous analysis. Nevertheless, selenium(IV) can be determined selectively and free of interferences by its reaction with NaBEt 4 [84]. In situ phenylation using sodium tetraphenyl borate has been studied with some applications [66]. Other reagents, such as sodium tetrapropylborate for organotin or tetramethylammonium tetrabutylborate for organolead [71], are promising but not readily available. Some efforts with multielement and multispecies derivatization using NaBEt4 [82] have been made. 2.3.4.3 Derivatizationwith Grignardreagents As mentioned above, hydride generation or ethylation with alkylborates fails for some species or in the cases of very complex matrices. An alternative is derivatization with Grignard reagents, which is versatile but requires an aqueous-free medium for the reaction to be carried out. In practice, derivatization with Grignard reagents is applicable to extracts containing complexes of an organometallic compound with dithizone, dithiocarbamates or tropolone. Whereas Grignard derivatization still remains the primary method for lead speciation analysis [49-51], its position [53] has been gradually eroding for organotin speciation in favor of the less cumbersome and time-consuming derivatization with NaBEt4 . Other applications of Grignard reagents, e.g., for the derivatization of organomercury, have been limited to one research group [57,58]. Grignard reagents proposed for derivatization in speciation analysis by GC with plasma source spectrometric detection have included: methyl-, ethyl-, propyl-, butyl- and pentylmagnesium chlorides or bromides. Lower-alkyl magnesium salts are generally preferred due to the smaller molecular mass and, hence, the higher volatility of the resulting species which makes the GC separation faster with less column carryover problems associated with derivatized inorganic forms (which are often present in large excess). In addition, the baseline is more stable and less Grignard reagent-related artifacts occur. Conversely, the low volatility of species derivatized by pentylmagnesium chloride may facilitate concentration by evaporation and, hence, a more efficient enrichment is achieved. 25

The unreacted Grignard reagent needs to be destroyed prior to the injection of the derivatized extract onto a column, which is achieved by shaking the organic phase with dilute H 2SO4 . As a final step of the procedure, the organic phase is dried over anhydrous Na2 SO4 , for example, and injected onto the GC column. 2.3.4.4 Other derivatization techniques The formation of volatile acetonates, trifluoroacetonates and dithiocarbamates is a popular derivatization technique for inorganic GC [147]. Kinetic restrictions or the thermodynamic inability of many species to react and the small differences in retention times for the derivatized species of the same element, make chelating agents of limited importance as derivatization reagents for speciation analysis. Selenoaminoacids were derivatized with isopropylchloroformate and bis(p-methoxyphenyl) selenoxide [148], with pyridine and ethyl chloroformate [149], or silylated with bis(trimethylsilyl)acetamide [150]. Selenomethionine forms volatile methylselenocyanide with CNBr [151,152]. Methods for the conversion of arsenic compounds to volatile and stable derivatives are based on the reaction of monomethylarsonic acid and dimethylarsenic acid with thioglycolic acid methyl ester (TGM) [153,154]. 2.3.5 Preconcentration and cleanup Some extraction techniques, such as sorption (solid-phase-extraction) or purge-and-trap offer a high intrinsic preconcentration factor. Solvent extraction methods have a common disadvantage of yielding a large volume of extract (usually about 5 ml) which increases considerably the amount that can be introduced onto the capillary column. The discrepancy between the concentration of an analyte species in a sample and the detector's sensitivity is often increased by a dilution factor during the analysis of solid samples. Therefore, an additional preconcentration step (e.g. by evaporation) sometimes needs to be carried out on the leachate or extract containing the analytes. Purging the extracts with nitrogen or helium in precalibrated tubes, Kuderna-Danish evaporation, and rotary evaporation have been the methods of choice. Losses may occur in the preconcentration of the derivatized species especially for more volatile Me3Pb+ species. Better recoveries are obtained when the solution of the organometallic chelates which are less volatile than tetraalkyl species is precon26

centrated prior to derivatization. Then, however, a minimum volume of 250 pl is required for easy handling during the derivatization step and removal of the unreacted Grignard reagent, introducing a dilution factor of 1:250 for capillary GC analysis. The increasing sensitivity of element selective detectors for gas chromatography (sub-picogram absolute detection limits become a standard) and the wider availability of large volume injection techniques have recently contributed to the elimination of the need for off-line enrichment. The evaporative preconcentration and large volume injection lead to a co-preconcentration of other substances present in a sample and may be detrimental to the chromatographic column in routine analysis. Therefore a cleanup step for the extract is advised. 2.3.6 Automation of sample preparation and GC sample introduction Figure 2.4, summarizing the recent advances in simplified and faster sample preparation and chromatographic (multicapillary) separation, 7 2 103

25

Alkaline digestion Acid digestion

1 95

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Sample preparation step 1 Digestion 2 []dilution 3 E]concentration

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Alkaline digestion Alkaline digestion

Microwave assisted

43

4

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7

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6 Elderivatization 7 centrifugation 8

Two-step procedure; ~conventional capillary chromatographic separation

One-step procedure; Conventional capillary chromatographic separation a One-step procedure; separation on a standard multicapillary column One-step procedure; separation on a custom designed multicapillary cartridge

o Elsample clean-up

|

0

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I

I

30

60

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I

120

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dry evaporation

9 OpH adjust

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11 1 phase separation 12 Eldetermination

I 180

210

Time, min Fig. 2.4. Simplification and acceleration of sample preparation protocols for speciation studies. 27

t

3

-way valve

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Drying gas o

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Fig. 2.5. Device for the time-resolved introduction of organometallic species into an atomic spectrometer.

indicates the possibility of development of an automated sample introduction device for an atomic spectrometer that would allow speciation analysis. Microwave-assisted leaching and derivatization seems to be a suitable technique for isolating the analytes from the sample matrix and adapting them to gas chromatography. Isothermal multicapillary GC seems to be the ideal separation technique to be incorporated in an automated speciation analyzer. A question remains regarding the interface between them (introduction of analytes onto a GC column). The automation of a system based on liquid-liquid extraction requires robotics and is cumbersome. A better perspective seems to be extraction of the liberated and derivatized analytes into the gas phase followed by their cryofocussing at the head of a GC column. The scheme of the accessory developed for time-resolved introduction of analytes into an atomic spectrometer is shown in Fig. 2.5. Organometallic compounds (alkyllead, butyltin, methylmercury) and some ions [Pb 2+ , Hg2 +, Se(IV), As(III) are in-situ volatilized by means of a suitable derivatization reaction (hydride generation or ethylation with NaBEt4 ). The derivatives formed are purged from the vessel and pass through a water scrubber (Nafion dryer) to a wide bore (0.53 mm) 28

capillary trap where they are cryofocussed at -100°C. Then, the trap is heated to release the species on a multicapillary capillary column. The analysis cycle takes less than 5 min. A number of applications have recently been described [155,156]. The system developed shows two major advantages over the commercial and the literature described purge-and-trap systems. This system is a free-standing accessory including the separation step, so no need for a gas chromatograph is required because of the application of a multicapillary column. The second advantage consists of using a 30 cm Nation tube dryer which makes an external chiller redundant. The result is a compact accessory allowing for species-selective analysis of liquid and solid (with optional microwave cavity) samples. 2.4 HANDLING OF BIOMATERIALS PRIOR TO HPLC A number of analytes cannot be volatilized and need to be separated by HPLC. The sample to be introduced on the column should be in an aqueous solution passing through a 0.2 pm filter. The major fields of interest have so far included speciation of organometalloid compounds of arsenic and selenium and speciation of metal complexes with bioligands (polypeptides and polysaccharides). Elemental speciation by liquid chromatography of compounds that can be determined by a GCbased coupled technique is of minor interest. 2.4.1 Organometalloid compounds The species of interest include As(III), As(V), mono- and dimethylarsenic acids, arsenobetaine, arsenocholine and a number of arsenosugars in biological materials. A freezing procedure has been commonly used to preserve a biosample; arsenic speciation in fresh and defrosted samples was carried out [157]. Arsenobetaine in sample extracts that were stored at 4C for 9 months was decomposed to trimethylarsine oxide and two other unidentified arsenic species [157]. Defatting, e.g. by leaching with acetone is the first step so as to avoid the generation of an emulsion with the fat, which would make the subsequent cleanup more difficult [158]. In addition, the efficiency of the subsequent methanol extraction step is apparently higher for defatted samples than for non defatted ones [158]. Extraction of arsenobetaine from tissue homogenates has usually been performed using methanol [158], methanol-chloroform-water or methanol-water. 29

With the BCR CRM 422, a precipitate of fatty appearance during CH 3OH-H 2 0 extraction was observed [159]. Chloroform is added in the classical procedure. Recoveries were 90% for fish and 80% for mussels [160]. No degradation of arsenobetaine to more toxic species was observed when an enzymic (trypsin) digestion procedure was applied to the fish [161]. After extraction with methanol, the supernatant is evaporated. The residue is suspended in water and shaken with diethylether to remove lipids [162]. A cleanup step was necessary due to an excess of lipids and fat in the extract [160]. Extraction with petroleum ether (5 times) was used [160]. Since some samples of seafood products are prepared in oil and generally tend to have a high salt content, an additional cleanup step, e.g. on strong cation exchanger (Dowex 50W-X8) [158] is required to eliminate the remains of liposoluble compounds not extracted with acetone. The clean-up also avoids the need for periodically reversing and flushing the chromatography column in order to overcome pressure buildup due to accumulation of material on the column. In contrast to organoarsenic, little is known on the identity of organoselenium compounds in biomaterials. The current studies of selenium speciation in yeast concerned either the water soluble fraction only (containing ca. 10% of all the selenium) [163-166] or aimed at the maximization of the Se recovery by degrading the species originally present with a mixture of proteolytic enzymes [167]. The results of the procedures have been contradictory. After leaching with an aqueous solution, Bird et al. [63,164] indicated the presence of more than 20 selenium compounds including selenocysteine, selenomethionine, methylselenocysteine, and inorganic forms present. The use of enzymatic hydrolysis lead Gilon et al. [167,168] to a conclusion that more than 80% of Se in the yeast was present in three forms: inorganic selenium, selenocysteine, and selenomethionine. Several procedures based on different principles were evaluated for recovery of selenium species from yeast [169]. Leaching with water and with methanol lead only to a 10-20% recovery of Se split into 8 compounds among which Se(IV) and selenomethionine could be identified. Leaching with pectinolytic enzymes released additionally 20% of selenomethionine. Leaching with a sodium dodecyl sulfonate solution allows the solubilization of a selenoprotein that accounted for ca. 30% of the total Se present. Leaching with proteolytic enzymes lead to recoveries of Se above 85%, the majority as selenomethionine. Hydrolysis of the yeast with tetramethylammonium hydroxide solubilized the 30

sample completely but the Se species present were entirely degraded to selenomethionine and inorganic selenium. 2.4.2 Coordination complexes of metals with bio-ligands 2.4.2.1 Biologicalfluids (full blood, plasma, synovial fluid, breast milk) Sample preparation of serum prior to HPLC involves filtration of sample on a 0.45 pm or 0.2 pm filter [170]. Erythrocytes were subjected to three freeze-thaw cycles to lyse the cells [171,172], followed by a tenfold dilution with a buffer and centrifugation at 18 000 g to remove fragments of membranes, etc. The supernatant was further diluted. Breast milk should be centrifuged to remove fat; precipitation of casein with 1 M acetate is optional [173]. Dialysis and purification by sizeexclusion chromatography is required if separation by RP HPLC should be undertaken [173]. 2.4.2.2 Plant and animal tissues Washing cells in a Tris-HCl buffer (pH 8) containing 1 M EDTA to remove metal ions reversibly bound to the cell wall was recommended [174]. In the majority of works size-exclusion chromatography (SEC) [175-178] has been preferred to heat treatment [179-181] for the isolation of the metallothionein (MT) fraction from the tissue cytosol. Guidelines for the preparation of biological samples prior to quantification of MTs were discussed with particular attention given to the care necessary to avoid oxidation [182]. Soluble extracts of tissues and cultured cells are prepared by homogenizing tissue samples in an appropriate buffer. Neutral buffers are necessary for extracting MTs since Zn starts to dissociate from the protein at pH 5. Cd and Cu are removed at lower pH values. A 10-50 mM Tris-HC1 buffer at pH 7.4-9 is the most common choice. For cytosols containing Cd-induced MTs, dilution factors up to 10 have been used, whereas for those with natural MT levels, equal amounts of tissue and buffer have been found suitable. Metallothioneins are prone to oxidation during isolation due to their high cystein content. During oxidation, disulphide bridges are formed and the MTs either copolymerize or combine with other proteins to move into the high molecular weight fraction. Since MTs may be oxidized by, e.g. oxygen, Cu(I) or heme components, the homogenization of tissues and subsequent isolation of MT should be normally performed in deoxygenated buffers and/or in the presence of a thiolic 31

reducing agent [183]. -Mercaptoethanol is added as an antioxidant that additionally prevents formation of dimeric forms of MT [184]. Other components added during homogenation include 0.02% NaN 3 which was added as an antibacterial and phenylmethanesulfonylfluoride which was added as a protease inhibitor. The homogenization step is followed by centrifugation. The use of an refrigerated ultracentrifuge (100 000 g) is strongly recommended. As a result, two fractions: a soluble one (cell supernatant, cytosol) and a particulate one (cell membranes and organelles) are obtained. Only the supernatant is analyzed for metallothioneins. Storage of the supernatant at -20°C under nitrogen prior to analysis is recommended [184]. Extraction efficiency by sucrose-Tris at pH 8 was 30-25% [182]. The percentage of metals in cytosols is typically 50-85% and 30-57% in the case of Zn and Cd, respectively [185,186]. A heat-treatment (at 60 ° for 15 min) of the cytosol extracts (especially of concentrated ones) is recommended to separate off the high-molecular fraction which coagulates from the supernatant (containing MTs which are heat stable). Such a treatment reduces the protein load on the HPLC column not only improving the separation of MT isoforms but also prolonging the column lifetime. Enzymolysis in simulated gastric and gastrointestinal juice was proposed for meat samples [181]. Filtration of the cytosol (0.22 pm filter) before introducing onto the chromatographic column is strongly advised. A guard column should be inserted to protect the analytical column particularly from the effects of lipids, which otherwise degrade the separation [181]. Any organic species that adhere to the column can also bind inorganic species giving rise to anomalous peaks in subsequent runs [181]. A new guard column was used for each injection to prevent adsorption by ligands with a high affinity for cadmium that would otherwise interfere with subsequent injections [181]. An extensive column cleanup was necessary [181]. To avoid contamination of the analytical column by trace elements, buffers should be cleaned by Chelex-100 [181].

2.5 CONCLUSIONS-TRENDS AND PERSPECTIVES Novel derivatization reagents and instrumental techniques have recently rendered the sample preparation prior to speciation of organometallic compounds by hyphenated techniques faster and easier to 32

automate. The ultimate goal to develop an automated speciation analyzer seems to be closer than ever. The critical issue remains the certainty that the species arriving at the detector from the chromatographic column is the one originally present in the sample. Controlling the stability of species for which standards can be synthesized, such as anthropogenic organometallic pollutants seems to be possible. Much, however, still remains to be done to control transformations of metal coordination complexes with bioligands during the sample preparation, especially since many of these species remain in complex chemical equilibria in a sample that is destroyed once a chemical or even a physical (dilution) operation is carried. Particular care is advised regarding the species stability during sampling and storage.

REFERENCES 1 G.E. Batley (Ed.), Trace Element Speciation: Analytical Methods and Problems. CRC Press, Boca Raton, 1987. 2 J.R. Kramer and H.E. Allen (Eds.), Metal Speciation:Theory, Analysis and Application. Lewis, Chelsea, 1988. 3 R.M. Harrison and S. Rapsomanikis (Eds.), Environmental Analysis Using Chromatography Interfaced with Atomic Spectroscopy. Horwood, Chichester, 1989. 4 P. Quevauviller, E. Meier and B. Griepink (Eds.), Method Validation in EnvironmentalAnalysis. Elsevier, 1995. 5 R. Lobinski, Appl. Spectrosc., 51 (1997) 260A. 6 P. Quevauviller, Method Performance Studies for Speciation Analysis. RSC, Cambridge, 1998. 7 R. Lobinski and Z. Marczenko, Spectrochemical Trace Analysis for Metals and Metalloids, Elsevier, Amsterdam, 1996. 8 C. Pecheyran, C.R. Quetel, F.M. Martin Lecuyer and O.F.X. Donard, Anal. Chem., 70 (1998) 2639. 9 J. Feldmann, J. Anal. Atom. Spectrom., 12 (1997) 1069. 10 J. Feldmann, R. Gruemping and A.V. Hirner, Fresenius'J. Anal. Chem., 350 (1994) 228. 11 J. Feldmann, and A.V. Hirner, Int. J. Environ. Anal. Chem., 60 (1995) 339. 12 W.H. Schroeder and R. Jackson, Chemosphere, 13 (1984) 1041. 13 T. Nielsen, H. Egsgaard, E. Larsen and G. Schroll, Anal. Chim. Acta, 124 (1981) 1. 14 R. Van Cleuvenbergen, W. Dirkx, P. Quevauvillier and F. Adams, Int. J. Environ. Anal. Chem., 47 (1992) 21. 33

15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 34

A.O. Valkirs, P.F. Seligman, G.J. Olson, F.E. Brincknman, C.L. Matthias and J.M. Bellama, Analyst, 112 (1987) 17. R.J. Carter, N.J. Turocy and A.M. Bond, Environ. Sci. Technol., 23 (1989) 615. D.E. Wells, Mikrochim. Acta, 109 (1992) 13. R. Ahmed and M. Stoeppler, Anal. Chim. Acta, 192 (1987) 109. P. Lansens, C. Meuleman, W. Baeyens, Anal. Chim. Acta, 229 (1990) 281. I. Heninger, M. Potin-Gautier, I. de-Gregori, H. Pinochet, Fresenius' J. Anal. Chem., 357 (1997) 600. M. Lucia and A.M. Campos, Mar. Chem., 57 (1997) 107. M.L. Mena, A. Morales-Rubio, A.G. Cox, C.W. McLeod and P. Quevauviller, Quim. Anal. (Barcelona), 14 (1995) 164. J.T. Van Elteren, J. Hoegee, E.E. Van der Hoek, H.A. Das, C.L. De Ligny, and J. Agterdenbos, J. Radioanal. Nucl. Chem., 154 (1991) 343. P. Quevauviller, M. Beatriz-de-la-Calle-Guntinas, E.A. Maier and C. Camara, Mikrochim. Acta, 118 (1995) 131. A.M. Ure, Quim. Anal. (Barcelona), 13 (1994) S64. J.L. Gomez-Ariza, E. Morales, R. Beltran, I. Giraldez, M. Ruiz-Benitez, Quim. Anal. (Barcelona]), 13 (1994) S76. P.J. Craig and P.D. Bartlett, Nature, 275 (1978) 635. F. Baldi, F. Parati and M. Filippelli, Appl. Environ. Microbiol., 59 (1993) 2479. P. Quevauviller, G.U. Fortunati, M. Filippelli, F. Baldi, M. Bianchi and H. Muntau, Appl. Organomet. Chem., 10 (1996) 537. R. Rubio and A.M. Ure, Int. J. Environ. Anal. Chem., 51 (1993) 205. J.F. Uthe and C.L. Chou, Sci. Tot. Environ., 71 (1988) 67. 0. Nygren and C. Nilsson, J. Anal. Atom. Spectrom., 2 (1987) 805. D. Chakraborti, W. Dirkx, R. Van Cleuvenbergen and F. Adams, Sci. Tot. Environ., 84 (1989) 249. S. Shawky and H. Emons, Chemosphere, 36 (1998) 523. R. Lobinski, Analusis, 22 (1994) 37. J. Szpunar, M. Ceulemans, R. Lobinski and F.C. Adams, Anal. Chim. Acta, 278 (1993) 99. M. Johansson, H. Emteborg, B. Glad, F. Reinholdsson and D.C. Baxter, Fresenius'J. Anal. Chem., 351 (1995) 461. H. Emteborg, D.C. Baxter, M. Sharp and W. Frech, Analyst, 120 (1995) 69. H. Emteborg, D.C. Baxter and W. Frech, Analyst, 118 (1993) 1007. P. Lansens, C. Meuleman, M. Leermakers and W. Baeyens, Anal. Chim. Acta, 234 (1990) 417. T. Perez-Corona, Y. Madrid-Albarran, C. Camara and E. Beceiro, Spectrochim. Acta, 53B (1998) 321. Z. Zhang, M.J. Yang and J.B. Pawliszyn, Anal. Chem., 66 (1994) 844A.

43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60 61 62 63 64 65 66 67 68 69

J.B. Pawliszyn, Solid-Phase Microextraction. Theory and Practice. Wiley VCH, Weinheim, Germany, 1997. T. Gorecki and J. Pawliszyn, Anal. Chem., 68 (1996) 3008. L. Moens, T. de Smaele, R. Dams, P. van den Broek and P. Sandra, Anal. Chem., 69 (1997) 1604. Y. Cai and J.M. Bayona, J. Chromatogr., 696 (1995) 113. Y. Morcillo, Y. Cai and J.M. Bayona, J. High. Res. Chromatogr., 18 (1995) 767. M. Radojevic, A. Allen, S. Rapsomanikis and M. Harrison, Anal. Chem., 58 (1986) 658. R. Lobinski and F.C. Adams, J. Anal. Atom. Spectrom., 7 (1992) 987. R. Lobinski and F.C. Adams, Anal. Chim. Acta, 262 (1992) 285. R. Lobinski, C.F. Boutron, J.P. Candelone, S. Hong, J. Szpunar-Lobinska and F.C. Adams, Anal. Chem., 65 (1993) 2510. M. Heisterkamp, T. de-Smaele, J.P. Candelone, L. Moens, R. Dams and F.C. Adams, J. Anal. Atom. Spectrom., 12 (1997) 1077. R. Lobinski, W.M.R. Dirkx, M. Ceulemans and F.C. Adams, Anal. Chem., 64 (1992) 159. J.A. Stab, W.P. Cofino, B. van Hattum and U.A.T. Brinkman, Fresenius'J. Anal. Chem., 347 (1993) 247. M. Ceulemans and F.C. Adams, Anal. Chim. Acta, 317 (1995) 161. K. Bergmann, U. Roehr and B. Neidhart, Fresenius'J. Anal. Chem., 349 (1994) 815. E. Bulska, H. Emteborg, D.C. Baxter, W. Frech, D. Ellingsen and Y. Thomassen, Analyst, 117 (1992) 657. D.C. Baxter and W. Frech, Anal. Chim. Acta, 12 (1991) 545. J.S. Blais and W.D. Marshall, J. Environ. Qual., 15 (1986) 255. D.S. Forsyth, D. Weber and C. Cleroux, FoodAddit. Contam., 9 (1992) 161. T.J. Gremm and F.H. Frimmel, Water Res., 26 (1992) 1163. H. Harino, M. Fukushima and M. Tanaka, Anal. Chim. Acta, 264 (1992) 91. J.L. Gomez-Ariza, E. Morales and M. Ruiz-Benitez, Analyst, 117 (1992) 641. M.J. Waldock, Mikrochim. Acta, 109 (1992) 23 T.M. Dowling and P.C. Uden, J. Chromatogr., 644 (1993) 153-160. V. Minganti, R. Capelli and R. De Pellegrini, Fresenius'J. Anal. Chem., 351 (1995) 471. M. Ceulemans, R. Lobinski, W.M.R. Dirkx and F.C. Adams, Fresenius'J. Anal. Chem., 347 (1993) 256. M. Ceulemans, J. Szpunar-Lobinska, W.M.R. Dirkx, R. Lobinski and F.C. Adams, Int. J. Environ. Anal. Chem., 52 (1993) 113. J. Szpunar, V.O. Schmitt, R. Lobinski and J.L. Monod, J. Anal. Atom. Spectrom., 11 (1996) 193. 35

S. Rapsomanikis, Analyst, 119 (1994) 1429. K. Bergmann and B. Neidhart, Fresenius'J. Anal. Chem., 356 (1996) 57. G.L. Hu, X.R. Wang, Y.R. Wang, X. Chen and L. Jia, Anal. Lett., 30 (1997) 2579. 73 N. Bloom, Can. J. Fish. Aquat. Sci., 46 (1989) 1131. 74 M. Horvat, N.S. Bloom and L. Liang, Anal. Chim. Acta, 281 (1993) 135. 75 M. Horvat, N.S. Bloom and L. Liang, Anal. Chim. Acta, 281 (1993) 153. 76 K.C. Bowles and S.C. Apte, Anal. Chem., 79, (1998) 395. 77 J. Dedina and D.L. Tsalev, Hydride-GenerationAtomic-Absorption Spectrometry. Wiley, Chichester, UK, 1996. 78 M. Filippelli, F. Baldi, F.E. Brinckman and G.J. Olson, Environ. Sci. Technol., 26 (1992) 1457. 79 K. Sato, M. Kohri, and H. Okochi, Bunseki Kagaku, 45 (1996) 575. 80 K. Jin, Y. Shibata and M. Morita, Anal. Chem., 63 (1991) 986. 81 S. Rapsomanikis, O.F.X. Donard and J.H. Weber, Anal. Chem., 58 (1986) 35. 82 M. Ceulemans and F.C. Adams, J. Anal. At. Spectrom., 11 (1996) 201. 83 P.J. Craig, R.J. Dewick and J.T. van Elteren, Fresenius'J. Anal. Chem., 351 (1995) 467. 84 B. de la Calle Guntifias, R. Lobinski and F.C. Adams, J. Anal. Atom. Spectrom., 10 (1995) 111. 85 Y. Liu, V. Lopez-Avila, M. Alcaraz and W.F. Beckert, J. High Resolut. Chromatogr., 17 (1994) 527. 86 S. Tutschku, S. Mothes and K. Dittrich, J. Chromatogr., 683 (1994) 269. 87 Y.K. Chau, F. Yang and R.J. Maguire, Anal. Chim. Acta, 320 (1996) 165. 88 E. Segovia-Garcia, J.I. Garcia-Alonso and A. Sanz-Medel, J. Mass Spectrom., 32 (1997) 542. 89 Y. Cai, S. Rapsomanikis and M.O. Andreae, Talanta, 41 (1994) 589. 90 Y. Cai, S. Rapsomanikis and M.O. Andreae, Anal. Chim. Acta, 274 (1993) 243. 91 G.L. Hu and X.R. Wang, Anal. Lett., 31 (1998) 1445. 92 Y. Cai, S. Rapsomanikis and M.O. Andreae, J. Anal. At. Spectrom., 8 (1993) 119. 93 F. Pannier, A. Astruc and M. Astruc, Anal. Chim. Acta, 287 (1994) 17. 94 A.M. Carro-Diaz, R.A. Lorenzo-Ferreira and R.J. Cela-Torrijos, J. Chromatogr., 683 (1994) 245. 95 H. Hintelmann, R.D. Evans and J.Y. Villeneuve, J. Anal. Atom. Spectrom., 10 (1995) 619. 96 R. Falter and G. Ilgen, Fresenius'J. Anal. Chem., 358 (1997) 407. 97 R. Falter and G. Ilgen, Fresenius'J. Anal. Chem., 358 (1997) 401. 98 R. Eiden, R. Falter, B. Augustin-Castro and H.F. Schoeler, Fresenius'J. Anal. Chem., 357 (1997) 439. 99 S. Padberg, M. Burow and M. Stoeppler, Fresenius'J. Anal. Chem., 346 (1993) 686. 70 71 72

36

100 N.S. Bloom, J.A. Colman and L. Barber, Fresenius'J. Anal. Chem., 358 (1997) 371. 101 S. Zhang, Y.K. Chau and A.S.Y. Chau, Appl. Organomet. Chem., 5 (1991) 431. 102 S. Hardy and P. Jones, J. Chromatogr.A., 791 (1997) 333. 103 R. Fischer, S. Rapsomanikis and M. Andreae,Anal. Chem., 65 (1993) 763. 104 S. Rapsomanikis and P.J. Craig, Anal. Chim. Acta, 248 (1991) 563. 105 M. Ceulemans, C. Witte, R. Lobinski and F.C. Adams, Appl. Organomet. Chem., 8 (1994) 451. 106 M.S. Jimenez and R.E. Sturgeon, J. Anal. At. Spectrom., 12 (1997) 597. 107 D.S. Forsyth and J.R. Iyengar, J. Assoc. Off. Anal. Chem., 72 (1989) 997. 108 D.S. Forsyth and J.R. Iyengar, Appl. Organomet. Chem., 3 (1989) 211. 109 F. Pannier, A. Astruc and M. Astruc, Anal. Chim. Acta, 327 (1996) 287. 110 Y. Liu, V. Lopez-Avila, M. Alcaraz and W.F. Beckert, J. High Resolut. Chromatogr., 16 (1993) 106. 111 C.M. Wai, Y. Lin, R. Brauer, S. Wang and W.F. Beckert, Talanta, 40 (1993) 1325. 112 Y. Liu, V. Lopez-Avila, M. Alcaraz and W.F. Beckert, Anal. Chem., 66 (1994) 3788. 113 J.M. Bayona and Y. Cai, Trends Anal. Chem., 13 (1994) 327. 114 W. Holak, J. Assoc. Off. Anal. Chem., 78 (1995) 1124. 115 R. Cela-Torrijos, M. Miguens-Rodriguez, A.M. Carro-Diaz and R.A. Lorenzo-Ferreira, J. Chromatogr. A., 750 (1996) 191. 116 I. Fernandez-Escobar and J.M. Bayona,Anal. Chim. Acta, 355 (1997) 269. 117 Y.K. Chau, F. Yang and M. Brown, Anal. Chim. Acta, 304 (1995) 85. 118 Y. Cai, R. Alzaga and J.M. Bayona, Anal. Chem., 66 (1994) 1161. 119 H. Emteborg, E. Bjorklund, F. Odman, L. Karlsson, L. Mathiasson, W. Frech and D.C. Baxter, Analyst, 121 (1996) 19. 120 B.E. Richter, B.A. Jones, J.L. Ezzel, N.L. Porter, N. Avdalovic and C. Pohl, Anal. Chem., 68 (1996) 1033. 121 F. Hoefler, LaborPraxis,20 (1996) 44. 122 J. Szpunar, V. Schmitt, O. Donard and R. Lobinski, Trends Anal. Chem., 15 (1996) 181. 123 O.F.X. Donard, B. Lalere, F. Martin and R. Lobinski, Anal. Chem., 67 (1995) 4250. 124 J. Szpunar, V.O. Schmitt, R. Lobinski and J.L. Monod, J. Anal. At. Spectrom., 11 (1996) 193. 125 J. Szpunar, M. Ceulemans, V. Schmitt, F. Adams and R. Lobinski, Anal. Chim. Acta, 332 (1996) 225. 126 I. Rodriguez Pereiro, V.O. Schmitt and R. Lobinski,Anal. Chem., 69 (1997) 4799. 127 I. Rodriguez Pereiro, A. Wasik and R. Lobinski, J. Chromatogr.A., 795 (1998) 359. 37

128 I. Rodriguez-Pereiro, A. Wasik and R. Lobinski, Chem. Anal. [Warsaw],42 (1997) 799. 129 C.M. Tseng, A. de Diego, F.M. Martin and O.F.X. Donard, J. Anal. At. Spectrom., 12 (1997) 629. 130 M.J. Vazquez, A.M. Carro, R.A. Lorenzo and R. Cela, Anal. Chem., 69 (1997) 221. 131 S. Shawky, H. Emons and H.W. Duerbeck, Anal. Commun., 33 (1996) 107. 132 I. Rodriguez, M. Santamarina, M.H. Bollain, M.C.Mejuto and R. Cela, J. Chromatogr. A, 774 (1997) 379. 133 C.M. Tseng, A. de-Diego, F.M. Martin, D. Amouroux and O.F.X. Donard, J. Anal. At. Spectrom., 12 (1997) 743. 134 I. Rodriguez, A. Wasik and R. Lobinski, J. Anal. At. Spectrom., 13, (1998) 743. 135 I. Rodriguez, A. Wasik and R. Lobinski, Anal. Chem., 70 (1998) 4063. 136 I. Rodriguez-Pereiro, V.O. Schmitt, J. Szpunar, O. Donard and R. Lobinski, Anal. Chem., 68 (1996) 4135. 137 I. Rodriguez-Pereiro and R. Lobinski, J. Anal. At. Spectrom., 12 (1997) 1381. 138 A.W. Kim, M.E. Foulkes, L. Ebdon, S.J. Hill, R.L. Patience, A.G. Barwise and S.J. Rowland, J. Anal. At. Spectrom., 7 (1992) 1147. 139 W.A. Aue, B. Miller and X. Sun, Anal. Chem., 62 (1990) 2453. 140 J.M. Ombaba and E.F. Barry, J. Chromatogr., 678 (1994) 319. 141 B.D. Quimby, P.C. Dryden and J.J. Sullivan, J. High Resolut. Chromatogr.,14 (1991) 110. 142 L. Ebdon, E.H. Evans, W.G. Pretorius and S.J. Rowland, J. Anal. At. Spectrom., 9 (1994) 939. 143 Y. Zeng and P.C. Uden, J. High Resolut. Chromatogr., 17 (1994) 217. 144 J.P. Snell, W. Frech and Y. Thomassen, Analyst, 121 (1996) 1055. 145 W. Frech, D.C. Baxter, B. Bakke, J. Snell and Y. Thomassen, Anal. Commun., 33 (1996) 7H. 146 H. Tao, T. Murakami, M. Tominaga and A. Miyazakin, J. Anal. At. Spectrom., 13 (1998) 1085. 147 G. Schwedt, ChromatographicMethods in Inorganic Analysis, Hithig, Heidelberg, 1981. 148 H. Kataoka, Y. Miyanaga and M. Makita, J. Chromatogr., 659 (1994) 481. 149 X.J. Cai, E. Block, P.C. Uden, X. Zhang, B.D. Quimby and J.J. Sullivan, J. Agric. Food Chem., 43 (1995) 1754. 150 K. Yasumoto, T. Suzuki and M. Yoshida, J. Agric. Food Chem., 36 (1988) 463. 151 Z. Ouyang, J. Wu and L. Xie, Anal. Biochem., 178 (1989) 77. 152 0. Zheng and J. Wu, Biomed. Chromatogr., 2 (1988) 258. 153 B. Beckermann, Anal. Chim. Acta, 135 (1982) 77. 154 H. Haraguchi and A. Takatsu, Spectrochim. Acta, 42B (1987) 235. 38

155 A. Wasik, I. Rodriguez and R. Lobinski, Spectrochim. Acta B, 53 (1998) 867. 156 A. Wasik, I. Rodriguez, C. Dietz, J. Szpunar and R. Lobinski, Anal. Commun., 35 (1998) 331. 157 S.X.C. Le, W.R. Cullen and K.J. Reimer, Environ. Sci. Technol., 28 (1994) 1598. 158 N. Ybanez, D. Velez, W. Tejedor and R. Montoro, J. Anal. At. Spectrom., 10 (1995) 459. 159 J. Alberti, R. Rubio and G. Rauret, Fresenius'J. Anal. Chem., 351 (1995) 415. 160 P. Thomas and K. Sniatecki, Fresenius'J. Anal. Chem., 351 (1995) 410. 161 S. Branch, L. Ebdon and P. O'Neill, J. Anal. At. Spectrom., 9 (1994) 33. 162 K. Shiomi, Y. Sugiyama, K. Shimakura and Y. Nagashima, Appl. Organomet. Chem., 9 (1995) 105. 163 S.M. Bird, P.C. Uden, J.F. Tyson, E. Block and E. Denoyer, J. Anal. At. Spectrom., 12 (1997) 785. 164 S.M. Bird, H. Ge, P.C. Uden, J.F. Tyson, E. Block and E. Denoyer, J. Chromatogr., 789 (1997) 349. 165 J. Zheng, W. GSssler and W. Kosmus, Trace Elem. Electrol., 15 (1998) 70. 166 H. Emteborg and G. Bordin, Analyst, 123 (1998) 245. 167 N. Gilon, A. Astruc, M. Astruc and M. Potin-Gautier, Appl. Organomet. Chem., 9 (1995) 623. 168 M. Potin-Gautier, N. Gilon, M. Astruc, I. De Gregori and H. Pinochet, Int. J. Environ. Anal. Chem., 67 (1997) 15. 169 C. Casiot, J. Szpunar, R. Lobinski and M. Potin-Gautier, J. Anal. At. Spectrom., 14 (1999) 645.. 170 G.F. Van Landeghem, P.C.D. D'Haese, L.V. Lamberts and M.E. De Broe, Anal. Chem., 66 (1994) 216. 171 I.A. Bergdahl, A. Schiitz and A. Grubb, J. Anal. At. Spectrom., 11 (1996) 735. 172 B. Gercken and R.M. Barnes, Anal. Chem., 63 (1991) 283. 173 Y. Makino and S. Nishimura, J. Chromatogr. Biomed. Appl., 579 (1992) 346. 174 K. Takatera, N. Osaki, H. Yamaguchi and T. Watanabe, Anal. Sci., 10 (1994) 567. 175 O.M. Steinebach and H.T. Wolterbeek, J. Chromatogr.Biomed. Appl., 130 (1993) 199. 176 M. Apostolova, P.R. Bontchev, C. Nachev and I. Sirakova, J. Chromatogr., B, Biomed. Appl., 131 (1993) 191. 177 S.G. Ang and V.W.T. Wong, J. Chromatogr., 599 (1992) 21. 178 H. Van Beek and A.J. Baars, At. Spectrosc., 11 (1990) 70. 179 A. Mazzucotelli and P. Rivaro, Microchem. J., 5 (1995) 231. 180 A. Mazzucotelli, A. Viarengo, L. Canesi, E. Ponzano and P. Rivaro, Analyst, 116 (1991) 605. 39

181 H.M. Crews, J.R. Dean, L. Ebdon and R.C. Massey, Analyst, 114 (1989, 895. 182 K.T. Suzuki and M Sato, Biomed. Res. Trace Elem., 6 (1995) 51. 183 K.A. High, B.A. Methven, J.W. McLaren, K.W.M. Siu, J. Wang, J.F. Klaverkamp, J.S. Blais, Fresenius'J. Anal. Chem., 351 (1995) 393. 184 H. Van Beek and A.J. Baars, J. Chromatogr., 442 (1988) 345. 185 C.K. Jayawickreme and A. Chatt, Biol. Trace Elem. Res., 503 (1990) 26. 186 K. Guenther and H. Waldner, Anal. Chim. Acta, 259 (1992) 165.

40