1848
Anal. Chem. 1086, 58, 1848-1852
(18) AnalaR Standards for hbwatory Chemicals, 6th ed.; Analar Standards, Ltd.: London, 1967; p 153. (19) Jones, M. H.; Woodcock, J. T. Anal. Chem. 1975, 4 7 , 11-16. (20) Jones, M. H.; Woodcock. J. T. Ultraviolet Spectrometry of Flotation Reagents with Special Reference to the Determination of Xanthate in
Flotation Liquors: The Institution of Mining and Metallurgy: London, 1973
(21)
Mingione, P.
A. In Ragants in the Minerals Industry; Jones, M. J., Oblatt. R., Eds.; The Instltution of Mlnlng and Metallurgy: London, 1984; pp 19-24.
for review February 18,1986. Accepted March 26,
1986.
Determination of Picogram Quantities of Methyltins in Sediment Cynthia C. Gilmour,' Jon H. Tuttle,*2 and J a y C. Means
Center for Environmental & Estuarine Studies and Department of Chemistry, Chesapeake Biological Laboratory, University of Maryland, Solomons, Maryland 20688-0038, and College Park, Maryland 20742
An extremely sensitlve purge and trap method is descrlbed for the determlnatlon of methyltins In complex matrlces. Organotlns were determined dlrectly from sediments and culture medlum as the volatlle methylstannanes. Hydrlde derivatives were prepared with NaBH, In a closed, flowthrough system conslstlng of a purge vessel, gas chromatograph, and mass spectrometer. Borate buffer added to samples generated H, from NaBH,, resulting In high purge efflclencles for mono-, dl-, and trhnethyttln. Selected lon mode monitoring with the masg spectrometer gave detectlon lknits for methyltins of 3-5 pg as Sn. The concentratlon detection llmits for a 5 3 sedlment sample were < w / g wfth a standard devlatlon of 6-18 "6, depending upon the methyltin specles and sample type. Sensitlvlty achieved was 2 orders of magnitude lower than previously reported for methyltins In sedlment. The method reported is both selective and specific, ellmlnatlng most Interferenceswhlle permitting podtlve ldentlflcatlon of lndlvldual methykin specles.
Methyltin species are ubiquitous in natural waters, although their concentration is usually low (less than 1 ng/L) in waters relatively unimpacted by anthropogenic activity (1,2). Monoand dimethyltin are the dominant species (1-3)) suggesting that methyltins, like methylmercury species, arise via stepwise methylation of the inorganic metal (4). Not only are sediment slurries capable of methylating added inorganic tin (5), but concentrtions of methyltin species increase with estuarine surface-to-volume ratios (1). Thus, tin methylation in aquatic environments likely occurs in sediments. Measurements of sediment methyltin concentrations show monomethyltin to be the dominant species in anoxic sediments while trimethyltin is found in highest concentrations in oxic sediments (6). This suggests that tin methylation probably occurs in anaerobic sediments, while degradation of higher molecular weight organotins such as tributyltin, an antifouling agent, occurs in oxygenated environments. In recent studies of inorganic tin methylation, we have confirmed that biomethylation occurs preferentially in anaerobic estuarine sediments (7). Methyltins were produced to a maximum level of about 2 ng/g (dry weight) of sediment in 21 days. Present address: Harvard University School of Public Health, Interdisciplinary Programs in Health, 665 Huntington Ave., Boston, MA 02115.
*Address correspondence to author a t Chesapeake Biological Laboratory, University of Maryland, Solomons, MD 20688-0038.
Methyltins may be extracted from complex matrices and analyzed by conventional gas-liquid chromatographic (GC) techniques (8). However, the procedure is lengthy, involving multiple steps where speciation may be altered and vessel adsorption effects may be large. Detection limits achievable with a flame ionization detector are 10-100 ng (9). Butylation of methyltin species before solvent extraction and use of atomic adsorption spectrophotometry shortens the extraction procedure and reduces detection limits to about 0.1 ng (IO). Analysis of organotins is often based on hydride generation, as the methylstannanes produced are both stable and volatile with boiling points ranging from 0 to 59 OC ( I , 2 , 6 , I I ) . Purge and trap (F/T) procedures followed by boiling point separations and detection by spectrophotometric methods yield detection limits in water of between 0.01 (2) and 1 ng (3). Detection of SnH emission by flame emission gives the greatest sensitivity (2). Chromatographic methods have also been applied with hydridization. Jackson et al. (12) used a commercial P/T apparatus fitted to a packed GC column and flame photometric detector to achieve a 0.1-ng detection. Our studies of microbial tin methylation required low part-per-trillion, or less than 10 pg, detection limits for methyltins in complex matrices such a~ bacterial culture medium and natural sediments. A rapid and direct method of analysis was preferable in order to preserve speciation and minimize transfer loss of tins on surfaces. We combined a modified hydride generation technique, used directly with sediment samples, with cryogenic trapping and selected ion mass spectrometry (SIM). The use of the P/T technique with SIM removes the majority of interferences while allowing positive identification and quantitation of methyltin species in sediment with detection limits of about 1 pg/g (dry weight) of sediment. Borate buffer added to samples increased the purging action of the hydridization reagent, NaBH4,by in situ production of H2 and allowed nearly complete recovery of methyltins added to 5 g wet weight of sulfidic sediment. This technique allowed measurement of part-per-trillion concentrations of methyltin species in a relatively pristine portion of Chesapeake Bay.
EXPERIMENTAL SECTION Apparatus. Methyltins were determined as their hydride derivatives with a Hewlett-Packard 5985B gas chromatograph (GC)-quadropolemass spectrometer (MS) equipped with a liquid nitrogen, cryogenic temperature programming accessory and an attached hydride generator. The hydride generator consisted of a 15 cm3glass readionlpurging vessel with side arm, screwed into a Teflon ring seal mounted in a stainless-steel headpiece. The headpiece was fitted with gas inlet and outlet lines. The hydride
0003-2700/86/0358-1848$01.50/00 1986 American Chemical Society
ANALYTICAL CHEMISTRY, VOL. 58, NO. 8, JULY 1986
1849
Table I. Response Curve Data for Methylstannane Species Calibrated against Diethyltin as Internal Standard
compd methyltin dimethyltin trimethyltin diethyltin
--
tR: min concn range, ng of
9.7 12.3
0.1-15 0.1-15
13.8
0.1-15
13.2
7
Sn N 6
6 6
slope, M (std dev)
R
intercept ( b )
4.174 (0.242) 4.076 (0.049) 4.841 (0.133)
0.991
0.501 0.161
0.999 0.995
0.024
detection limit, pg of Sn 4.0 3.0 4.7
tR = retention time.
genertor was connected directly to the GC carrier gas supply via a zero-dead-volume, six-port stainless-steel valve (Valco). The GC column consisted of a 2-mm4.d. X 2-m glass column packed with 0.2% Carbowax 1500 on 80/100 mesh Carbopak C (Supelco). Zero-grade He at a flow of 30 mL/min served as both the purging gas and the carrier gas. A glass jet separator (Scientific Glass Engineering) passed column eluent to the MS. The MS was operated in electron impact mode at an electron energy of 70 eV. Typical MS operating conditions were as follows: ion source pressure, 3 x 10-6 torr; source temperature 200 "C; electron multiplier voltage, 2400 V. Comparative determinations were performed with a Hewlett-Packard 7676A purge and trap sampler containing a Tenax-GC filled 0.5 X 10 cm2stainless-steeltrap. This sampler was used with a GC column copacked with 3% SP 2401 and 10% SP2100 on 80/100 mesh Supelcoport. Reagents. Mono-, di-, and trimethyltin chlorides and diethyltin dichloridewere obtained from Alfa Chemicals. Standard solutions (10 mg mL-') were made in glass-distilled methanol (Burdick and Jackson) and stored at -10 "C. These solutions were stable for at least 6 months. Working dilutions were prepared daily in water deionized with a Milli-Qwater system (Millipore)and then glass distilled. Working standards were kept ice-cold during analysis. Reagent grade chemicals and deionized-distilledwater were used to prepare the derivitization solution (4% NaBH4,1% NaOH in water) and dilution buffer (saturated sodium borate, adjusted to pH 8.0 with NaOH). Procedure. Methylstannanes were generated directly in buffered sample, purged, and cyrogenicallytrapped on the head of a chromatographic column. Gas flow was diverted from the hydride generator while the purge tube was filled with up to 10 mL of fluid, usually consisting of 5 mL of sample and 5 mL of buffer. The purge tube was then sealed to the Teflon head. Samples were spiked with known amounts of diethyltin dichloride, which served as internal standard. The GC column was held at -40 "C for 2 min before 1 mL of NaBH4 solution was injected into the purge vessel through a butyl rubber septum on the vessel sidearm. Helium flow was then directed through the hydride generator for 3 min. After the purge cycle was completed, carrier gas was again diverted away from the sample and the column oven temperature then increased at 15 "C/min to 120 "C, then 30 "C/min to 150 "C. MS operation began 6 rnin after the purge cycle was complete and operated 7.5 min. Identification and quantitation of methyltin hydrides were achieved by selected ion monitoring (13). Major ions representative of the three methylstannanes and the internal standard (diethyltin) were nominal, m / e 116, 118,120, 133, 135, 148, 150, and 165. Sediment Sampling. Sediments were collected from various sites in Chesapeake Bay with a Van Veen grab or with 5-cm-i.d. Lexan core tubes. Grab samples were held in sealed 40-L containers until use. Core samples were immediately subsampled by removing minicores that were then frozen. Minicores were removed at 2.5-cm depth intervals from the larger cores by inserting cut-off 10 cm3polyethylene syringes into the whole cores through appropriately sized holes and sealing the open end of the syringe with butyl rubber serum stoppers. All glass and plasticware was presoaked in 10% "0% Buffer stored for 1 week in a sealed syringe showed no detectable methyltins upon analysis. RESULTS AND DISCUSSION Calibration, Precision, a n d Detection Limits. Response curve data for mono-, di-, and trimethyltin obtained with the
/DMT 41
Methyltin Concentration, ng Figure 1. Response curves for mono- (O),di- (A),and trimethyltin (0) calibrated with 7 ng of diethyltin as internal standard.
cryogenic trapping GC-MS method were based on the response of diethyltin internal standard (Figure 1) . Figure 2 shows typical selected ion chromatographs of mixtures of the three methyltins and internal standard. Retention times of the methylstannanes depended on their boiling points, as described by Hodge et al. (3). Table I lists least-squares analyses of the response factor data for the three compounds over a range of slightly more than 2 orders of magnitude in concentration. All three methylstannanes gave linear response curves over the range tested. Absolute detection limits were calculated by solving the calibration regression equations for twice background noise. For typical sample volumes of 5 mL of bacterial culture medium or 1-3 g (dry weight) of sediment, the detection limits were equivalent to concentrations of about 0.9 pg of Sn/mL or between 1.3 and 5 pg of Sn/g of sediment (dry weight). Purge Efficiency. The total volume purged was routinely adjusted to 10 mL with borate buffer. Boric acid and Na13H4 react to produce hydrogen gas, increasing the efficiency of methylstannane purging over that produced by helium gas flow alone. This is especially useful in sediment analyses, as the hydrogen moves through a larger portion of the interparticle spaces than the portion scrubbed by a single stream of purge gas. At a He flow rate of 30 mL/min, no increase in peak areas occurred as purge times were increased past 3 min. A second analysis of an already purged sample yielded no detectable organotin compounds. We also observed no decrease in peak area with increasing purge time, and no tin species were detected when the MS was operated during the purge cycle. This indicates that volatile methylstannanes, especially monomethylstannane with a boiling point of 0 "C,
1850
ANALYTICAL CHEMISTRY, VOL. 58, NO. 8, JULY 1986
1
TMT
l MMT A ~
DMT
I
A
m
fn 0
P
m
U
B
II
8
10
12
Retention Time (minutes) Flgure 3. Comparative selected ion chromatograms of 50 ng each of mono-, di-, and trimethyltin (A) and 50 ng of monomethyttin only (6) trapped on Tenax-GC and separated at 30 O C on a GC column packed with 3% SP2401 and 1 0 % SP-2100 on 80f100 mesh Supelcoport. Retention Time (minutes)
Flgure 2. Selected ion chromatograms of methylstannanes generated from a d d i i of 10 ng (A) or 0.1 ng (6)each of mono- (MMT), dL (DMT), and trimethykln (TMT) containing 7 ng of dlethyltrn (DET) as internal standard in borate buffer or 10 ng (C) of each methyltin and 7 ng of DET added to anoxic, sulfidic sediment from Chesapeake Bay.
did not break through the cryogenic GC trap during the purge cycle. The use of a commercial purge and trap apparatus (Hewlett-Packard 7675A P/T GC attachment) in the analysis of methylstannanes was also explored. This system contains a metal 1-cm-i.d.trap packed with Tenax-GC, a molecular seive type packing. Organotins were trapped at room temperature and desorbed onto a packed GC column by ballistically heating the trap to at least 200 "C. Detection limits with this method were higher than for the cryogenic trapping method, about 20 pg for di- and trimethyltin. They were especially poor for monomethyltin, which had an absolute detection limit of about 0.2 ng. These results are comparable to the use of this P/T apparatus with a flame emission detector (12). We found that monomethylstannane began to break through the trap before trimethylstannane purging was complete. There was also evidence of rearrangement of monomethylstannane on the Tenax trap, yielding a significant peak at the retention time of dimethylstannane, which was not found in blanks (Figure 3B). This peak gave a selected ion spectrum identical with that of dimethylstannane. Peak shapes were also broader than during cryogenic trapping. However, chromatographic separation used with the Tenax-GC trap was carried out a t 30 "C, and peak shapes would likely improve if separation was done at lower temperatures. Recovery of Methyltins Added to Sediment. The recovery of methyltins from sediments was tested by using anoxic, sulfidic clay sediments from a mid-salinity region of Chesapeake Bay and using liquid cultures of sulfate-reducing bacteria. The medium for growth of sulfate reducers contained complex organic compounds such as vitamins, as well as simple sugars and salts at levels comparable to or higher than those found in the estuarine environment. The cultures also contained bacterial cells (about lo9 cells/mL) and their products, including millimolar levels of sulfide. Mono- and dimethyltin were completely recovered from sediment (Table 11). However, recovery of 10 ng of trimethyltin chloride from sediment was only about 70% under the purging conditions described above. The percent recovery of both di- and trimethyltin from
Table 11. Recovery of 10 ng Each of Mono-(MMT), Di-(DMT), and Trimethyltin (TMT) Standards Added to Sediment or Bacterial Cultures
MMT
sediment
av % recovery (%
std dev)
culture
10.17 10.81 7.30 11.19
10.39 11.58
8.85
6.75 7.09 6.01
11.49
98.7
105.8
69.1
(17.6)
(12.7)
(7.3)
10.04 9.78 11.24 8.94
7.00 7.01 7.99 6.61
5.51 6.64 7.34 4.76
av % recovery
60.6
std dev)
(11.5)
(%
amt recovered, ng DMT TMT
100.0 (9.5)
71.5 (5.9)
bacterial culture was similar to that from sediment, although the monomethyltin recovery from cultures was lower, about 60%. Increased purge time might improve recovery of trimethyltin. However, as monomethylstannane is the most volatile of the three tested methylstannanes, low recovery of monomethyltin in bacterial culture is likely due to chemisorption effects and would not be improved by increased purging. Although little information is available concerning the binding constants of methyltins to the components of natural waters, precipitates from unfiltered seawater spiked with mono- and dimethyltin contained all the added monomethyltin while all of the dimethyltin remained in solution (3). Reported values for monomethyltin in microbial culture media must therefore be considered as minimum estimates. Methyltin Determination in Environmental Samples. Sediments from three sites along a mid-salinity transect of Chesapeake Bay were analyzed for methyltin content as a function of depth. These sediments, expected to be low in methyltins relative to contaminated harbor and river sites, were chosen so that the method could be evaluated for use in complex sulfidic samples containing low methyltin levels. The transect extended between the Patuxent River, MD, on the western shore and Barren Island on the eastern shore. Sediments at all three sites were anoxic and sulfidic below 1 cm depth. the eastern-most site, in the shipping channel
ANALYTICAL CHEMISTRY, VOL. 58, NO. 8, JULY 1986
1 1
1851
A
RT- 9.7 min
DET
Retention Time (minutes)
Flgwe 4. Selected ion chromatograms of methylstannanes generated from Chesapeake Bay sediments with 7 ng of DET as tntemal stanbard: (A) sediment from shipping channel and (B) nearshore sediment.
with a water depth of 44 m, had black, silty clay sediments. An intermediate site, with a depth of 14 m, also had black clay sediments. The third site was located on the shallow western shelf in 7.5 m of water. Sediments here were silty-sand and greatly bioturbated. Preweighed subsamples of minicores taken at 2.5-cm depth intervals from a larger core at each site were spiked with diethyltin internal standard, mixed with 5 mL of borate buffer, and analyzed directly. Other subsamples were used for sediment dry weight determination. Selected ion chromatograms of the methylstannenes evolved from two of these sediments are shown in Figure 4. Monoand dimethyltin were found a t every depth examined a t all three sites. Trimethyltin was frequently detectable, though not in every sample. Tin species were initially identified by retention time and confirmed by the selected ion spectrum of each peak. Selected ion spectra for monomethyltin from a known standard and from sediment are given in Figure 5A,B. The chromatogram of nearshore sediment (Figure 4) shows peaks a t retention times other than those identified with methylstannanes. The selected ion spectrum (Figure 5C) of the largest of these peaks (retention time 12.8 min, Figure 4B) shows ion ratios incompatible with the natural ratios of tin isotopes, confirming that the peak does not represent a tin compound. The ability to positively identify eluted compounds by their partial mass spectra significantly increases the selectivity of this method over spectrophotometric determinations. Methyltin concentrations in sediments along the mid-bay transect were quite low. Concentrations of all three methyltin species generally decreased with depth. Monomethyltin was predominant in most samples, with maximum concentrations of 0.18 and 0.13 ng/g (dry weight) for the shallow and deep sites, respectively. At the middle site all species were present at an average of less than 5 pg/g (dry weight). We were not able to compare methyltin concentrationsin anoxic Sediments to levels in oxic sediments, as all cores collected were anaerobic below 1cm depth, and thus all subsamples were taken from the anoxic zone. Sediment samples from Baltimore Harbor and Solomons Harbor, on the Patuxent River, MD, were also analyzed. Both sediments were anoxic with high sulfide levels. Baltimore Harbor sediments averaged 8 ng of mono-, 1ng of di-, and 0.3 ng of trimethyltin/g dry weight of sediment. Methyltin concentrations in relatively unpolluted Patuxent sediments were much lower, averaging about 1 ng of mono-, 0.1 ng of di-, and 0.01 ng of trimethyltin/g dry weight. Tugrul et al. (6)measured the concentration (0.6 ng/g dry weight) of monomethyltin in sediment from an unidentified location in
r
801
II C
R T - 12.8 min 1
I
I
401
t 120 L 4130d 140 L - 150 L
160
mle
Figure 5. Selected ion spectra of methyistannane species: (A) monomethylstannane generated from standard in borate buffer, (B) monomethylstannane evolved from nearshore sediment, (C) spectrum of
non-tin compound generated from nearshore sediment. Chesapeake Bay by extraction, hydridization, cryogenic trapping, and atomic absorption (AA) detection. Their value falls just above values for mid-bay sediments,just below values for Patuxent River sediments, and about an order of magnitude below Baltimore Harbor values given here. Di- and trimethyltin were not detectable in the sediment analyzed by the AA method (6). The cyrogenic GC-MS method represents an improvement in sensitivity of at least 2 orders of magnitude over the only other previous analysis method for methyltins in sediments, i.e., the method of Tugrul et al., which involves multiple steps including extraction (6). Our sediment method,is equal in detection limit and linear range to the most sensitive method described for analysis of methyltins in water, a similar purge and trap technique used with flame emission detection (2). Selective purging of methylstannanesfrom samples and using MS for detection eliminate nearly all interferences and stannanes are positively identified. The method especially eliminates a major interference from sulfur compounds that occurs when FPD is utilized (12). This became quite important, as we identified sulfidic environments and the sulfate-reducing bacteria contained therein as the likely producers of methyltins in the estuary (7). Direct analysis of sediment samples at near in situ pH (8.0) without pretreatment decreases transfer losses and the possibility of changes in speciation. Due to the sensitivity obtainable, care must be taken to eliminate interference due to leaching of organotin plasticizers from poly(viny1 chloride) pipes and containers, and from some plastic labware. Near total recovery of methyltin species from sediment, combined with picogram detection limits, allows measurement of methyltin species at pristine sites. It has also allowed the study of the relatively slow time course of microbial methylation of inorganic tin a t natural levels, both in bacterial culture and in natural sediments (7).
ACKNOWLEDGMENT We thank F. Brinckman and G. Olson of the National Bureau of Standards, Gaithersburg, MD, for use of their HP 7675A P/T sampler.
1852
Anal. Chem. 1986, 58, 1852-1857
Registry No. MMT, 23001-26-5; DMT, 23120-99-2; TMT, 1631-73-8; DET, 660-74-2.
LITERATURE CITED (1) Byrd, J. T.; Andreae, M. 0. Scknce (Washlngton, D.C.)1982, 278, 565-569. (2) Braman, R. S.;Tompkins, M. A. Anal. Chem. 1979, 5 7 , 12-20. (3) Hodge, V. F.; SeMel, S.L.; Goldberg, E. D. Anal. Chem. 1979, 5 1 , 1256-1259. (4) Ridley, W. P.; dizlkes, L. J.; Wood, J. M. Science (Washington, D.C.) 1977, 197, 329-332. (5) Hallas, L.E.; Means, J. C.; Cooney, J. J. Sclence (Washington, D.C.) 1982, 275, 1505-1506. (6) Tugrul, S.; Balkas, T. I.; Goldberg, E. Mer. PONot. Bull. 1983, 1 4 , 297-303. (7) GHmour, C. C.; T W , J. H.; Means, J. C. I n Merlne and Estuarlne Geochemlsby; Slgleo, A. C.; Hattori, A; Eds.; Lewis Publishers, Chelsea, MI. 1985; pp 239-258. (8) Arakawa, Y.; Wada, 0.; Yu, T. H.;Iwai, H. J. Chromatogr. 1981, 216, 209-217.
(9) Tam, 0. K.; Lacrolx, G.; Lawrence, J. F. J. Chromatogr. 1983, 259, 350-352. (10) Chau, Y. K.; Wong, P. T. S.; Bengert, G. A. Anal. Chem. 1982, 54 246-249. (11) Andreae, M. 0.; Byrd, J. T. Anal. Chim. Acta 1984, 756, 147-157. (12) Jackson, J. A.; Blair, W. R.; Brlnckman, F. E.;Iverson, W. P. Environ. Sci. Techno/. 1982, 76, 110-119. (13) Means. J. C.; Hulebak, K. L. Neurotoxicology 1983, 4 , 37-44.
RECEIVED for review November 4,1985. Accepted March 17, 1986. This work was supported in part by Grant OCE 8208032 from the National Science Foundation. C.C.G. was supported by a Chesapeake Biological Laboratory graduate research assistantship. Contribution No. 1683 of the Center for Environmental & Estuarine Studies of the University of Maryland.
Screening of Anthropogenic Compounds in Polluted Sediments and Soils by Flash EvaporationlPyrolysis Gas Chromatography-Mass Spectrometry J. W. de Leeuw,* E. W. B. de Leer, J. S . Sinninghe Damst6, and P. J. W. Schuyl Department of Chemistry and Chemical Engineering, Delft University of Technology, de Vries van Heystplantsoen 2, 2628 RZ Delft, The Netherlands
The use of flash evaporation and pyrolyrb gas duomstography-mass spectrometry as a fast screenbg procedwe for anthropogenic suWancw In envkoMnental samples b demonstrated by the analyrk 0t poUuted sol1 and 8edlmont samples. Pdycyck aromstk hydrocarbons, haloorganla, allphatlc hydrocarbons, heteroaromatics, elemental wlfur, cyanides, and pyrdyrk products of synthetk polymers are among the anthropogenic substances that can be readlly detected by this method In one analysis. ElLnkratbn of wet chemkal uimplepr.paraUon enables a complete analyrio to beperfan\.danddatatobequlddyMdyzed. Thedetectkn lknltrr are In the low part-per-mlllkn range wlng mass spectrometric detectlon. Alteratlvely, detectlon of compounds can be achleved by all common gas chromatography decectm (flame knltstkndetector, docban captwe detector, and flame photometrk detector), and detectkn lknits are determined by the method of detection employed.
Qualitative and quantitative analysis of pollutants in soils and sediments is an expensive and time-consuming task. Moreover, each suite of pollutants normally requires other analytical methods and techniques, so a large number of separate analyses have to be performed before a more or less complete picture of the pollution pattern is obtained. Recently two different fast screeningprocedures to monitor organic contaminanta in the environment have been reported (1, 2). Hunt et al. (1) used a triple quadrupole mass spectrometer configuration to analyze various series of contaminants with few wet chemical and chromatographic separation steps. McMurtrey et al. (2) reported a fast procedure that also omits the usual extraction and cleanup steps to monitor 0003-2700/86/0358-1852$01 SO10
the presence of polychlorinated biphenyl contaminants in a sediment using a pyrolysis gas chromatography-mass spectrometric technique. Over the last years we have used similar pyrolysis techniques, in particular Curie point flash pyrolysis mass spectrometry (Py MS) and Curie point flash pyrolysis gas chromatography-mass spectrometry (Py GC-MS), to chemically characterize biopolymers such as lignins, polysaccharides, peptides, and “geopolymers” like humic substances, peak, kerogens, and coal (3-10). During these studies it became clear that nonpolymeric compounds which are reasonably volatile at elevated temperatures do not fragment on the pyrolysis wire but simply evaporate from it (10-12). Therefore, it appeared possible that the organic matter present in sediments and soils can be characterized and identified very rapidly without any pretreatment by direct evaporation/pyrolysis (Ev/Py) of whole samples. This flash evaporation/pyrolysis method followed by on-line separation and identification techniques is thought to be applicable to monitor the volatile and nonvolatile (polymeric) contaminants in environmental samples such as soils, sediments, and tissues. In this paper we demonstrate the usefulness of Ev/Py GC-MS as a screening technique for anthropogenic substances in two polluted environmental samples without pretreatment. The samples were also extracted, and some compounds present in the extract were quantitated to establish the limits of detection of the screening approach.
EXPERIMENTAL SECTION Samples. Polluted soil (1m2)was removed with a spade from the top 10 cm from a site in the northwestern part of the Netherlands and thoroughly mixed. Approximately 2 kg of this wet soil was placed in a stainless-steel jar and brought to the lab. Samples were further homogenized by grinding before analysis. 0 1986 American Chemical Society
