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Identification of dimethylchloroarsine near a former herbicide factory by headspace solid-phase microextraction gas chromatography-mass spectrometry
Chemosphere 48 (2002) 1003–1008 www.elsevier.com/locate/chemosphere
Identification of dimethylchloroarsine near a former herbicide factory by headspace solid-phase microextraction gas chromatography-mass spectrometry Daniel R. Killelea 1, Joseph H. Aldstadt, III
*
Department of Chemistry, University of Wisconsin-Milwaukee, 3210 North Cramer Street, Milwaukee, WI 53211, USA Received 7 December 2000; received in revised form 27 March 2002; accepted 30 March 2002
Abstract The application of an improved method based on multidetector gas chromatography to the determination of trace levels of organoarsines in complex matrices is described. The method using headspace-mode solid-phase microextraction (SPME) was applied to a carefully sampled and preserved freshwater sediment core obtained from central Green Bay, Lake Michigan. The sediment core was collected and fractionated in an inert atmosphere. A carboxen/ polydimethylsiloxane-coated SPME fiber (85 lm film thickness) was equilibrated (n ¼ 4) for 60 min at 25 °C in the headspace of the sample vessel before introduction to the chromatograph. Conventional quadrupole ion trap mass spectrometry (electron impact ionization), electron capture detection, and pulsed flame photometric detection (arsenic mode) were employed for structure elucidation. A heretofore unidentified species in this region, dimethylchloroarsine (DMCA), was identified. The mass spectrum for DMCA is interpreted based on the observed fragmentation pattern. A bimodal vertical distribution of DMCA in the sediment core sample was observed and its interpretation based on Pb210 dating is reported. Ó 2002 Published by Elsevier Science Ltd. Keywords: Arsenic; Green bay; Dimethylchloroarsine [CAS registry no. 557-89-1]
1. Introduction Arsenic is ubiquitous in nature, and it is now firmly established that the impact of environmental arsenic cannot be expressed simply as a function of total arsenic concentration (Ferguson and Gavis, 1972). Indeed, the many compounds that this reactive metalloid can form encompass a wide range of physicochemical and toxicological properties (Matschullat, 2000). To better understand transport and ultimate effect on a given ecosystem, it is critical to identify and quantify all of the specific
*
Corresponding authors. Tel.: +414-229-5605; fax: +414229-5530. E-mail address: [email protected] (J.H. Aldstadt, III). 1 Present address: Department of Chemistry, Tufts University, 62 Talbot Avenue, Medford, MA 02155, USA.
chemical forms of arsenic that are present (Cullen and Reimer, 1989). Providing accurate information on the abundance, distribution, and structures of arsenicals at environmentally relevant levels is analytically challenging. The chemical species of arsenic in the environment are typically divided into five general classes: (a) inorganic covalent oxygen and sulfur forms, such as As2 S3 and As2 O3 ; (b) the inorganic protonated oxyanions, i.e., arsenious acid, As(OH)3 , and arsenic acid, AsO(OH)3 ; (c) alkylated derivatives of the acids, including, monomethylarsonic acid (MMAA), dimethylarsinic acid (DMAA), trimethyl arsine oxide, arsenobetaine, and arsenocholine; (d) the so-called ‘‘arsenosugars’’, with most attention focusing on the major species found in marine organisms; and (e) the arsines (Penrose, 1974; Andreae, 1977; Andreae, 1979; Anderson and Bruland, 1991).
0045-6535/02/$ - see front matter Ó 2002 Published by Elsevier Science Ltd. PII: S 0 0 4 5 - 6 5 3 5 ( 0 2 ) 0 0 1 6 0 - 1
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D.R. Killelea, J.H. Aldstadt, III / Chemosphere 48 (2002) 1003–1008
The major forms of arsenic that fall into the ‘‘arsine’’ category are inorganic arsine (AsH3 ) and aliphatic (methylated) derivatives of trivalent arsenic––monomethyl arsine (MMA), dimethyl arsine (DMA), and trimethyl arsine). Arsines can be generated by bacterial or fungal action, e.g., the classic example is the action of the fungus Scopulariopsis brevicaulis in producing trimethylarsine (Challenger et al., 1933; Andreae, 1986). As a class, the arsines represent some of the most toxic forms of arsenic, and under certain conditions can be easily transported in nature because of their pronounced volatility. At the same time, the arsines are generally reactive toward hydrolysis and oxidation, thus in many instances mitigating their potentially detrimental effects. While methods for measuring total arsenic and most major arsenic species are numerous (Irgolic, 1992), there are relatively few modern methods for analysis of the arsines in environmental samples (Le and Ma, 1998; Pantsar-Kallio and Korpela, 2000). Equally importantly, there is substantial inconsistency in the procedures used to collect arsenicals in the field in general and arsines in particular prior to their determination in the laboratory (Cullen and Reimer, 1989). The difficulty in measuring the arsines usually arises in the sampling phase of analysis because one must exercise great care to accurately sample such volatile and potentially reactive compounds. Recently we described a method for the determination of several organoarsenic acids that combined chemical derivatization with solid-phase microextraction (SPME) and gas chromatography-mass spectrometry (GC-MS) (Szostek and Aldstadt, 1998; Killelea and Aldstadt, 2001). For polar and/or thermallylabile analytes, analyte derivatization prior to SPME-GC can enhance detectability and improve chromatographic resolution (Pawliszyn, 1997). We examined a series of dithiol compounds for derivatization of the arsenicals, and the best results were obtained with 1,3-propanedithiol (PDT). The derivatization procedure, fiber type, and extraction time were then optimized. Using this method, the limit of detection for 2-chlorovinyl arsonous acid was improved 400 fold compared to conventional solvent extraction methods. The goal of the work reported herein was to modify our previous SPME GC-MS method (Killelea and Aldstadt, 1998) by using headspace-mode SPME without thiol derivatization to carefully preserved freshwater sediments near a former herbicide manufacturing facility to determine if organoarsines were present.
prepared using a NanoPureTM filtration system equipped with an ultraviolet lamp (Barnstead-Thermolyne, Dubuque, IA, USA). Arsenic standards were obtained from Aldrich Chemical (Milwaukee, WI, USA). All glassware and plasticware were acid-washed after use for at least 48 h in 5% (v/v) AR-grade nitric acid to remove background contamination. Arsenic-containing standards with concentrations less than 1 mg/l were prepared on the day of use. All aqueous standards were stored at 4 °C. 2.2. Instrumentation Organoarsenicals were determined by gas–liquid chromatography using a Varian GC-MS system (Walnut Creek, CA, USA) located in a Class 100 clean room. The GC-MS consisted of the following components: Model 3800 capillary gas–liquid chromatograph (DB5MS column, 30 m 0:25 mm with 0.25 lm film, J&W Scientific, Folsom, CA, USA) with Model 1079 split/ splitless injector; SPME apparatus (Supelco, Bellefonte, PA, USA); electron impact ionization source (70 eV); Saturn 2000 quadrupole ion trap MS (10–650 m=z range, unit resolution). The automatic gain control of the MS system was used throughout this study. Automated library searching was performed using the NIST Mass Spectral Database (version 3.0). The mobile phase was ultrahigh purity (99.999%) helium (Praxair, Milwaukee, WI, USA) at a constant linear velocity of 42 cm/s by electronic flow control. Samples were also studied using electron capture detection and pulsed flame photometric detection, as described elsewhere (Killelea and Aldstadt, 2001). 2.3. GC program For the SPME work, the injection port was 250 °C for the fiber desorption step (splitless). The initial column temperature was 45 °C for 5 min, then programmed at 20 °C/min to 160 °C, followed by 8 °C/min to 210 °C and finally 50 °C/min to 300 °C, where it was held for 5 min. The transfer line between the GC and MS was maintained at 170°. Mass spectra were obtained by scanning from m=z 35 to 400 with a 0.7 s scan time. GC instrument control and data acquisition were performed on a Pentium personal computer (Dell, Optiplex GX1, Dallas, TX, USA) using Saturn System software version 5.21 (Varian).
2. Experimental
2.4. Sediment procedures
2.1. Reagents
Sediment samples were collected in July 1999 aboard R/V Neeskay at a depth of approximately 20 m in central Green Bay, approximately 13.5 km south of the mouth of the Menominee River (Green Bay Station 40).
All chemical reagents used were analytical reagent (AR) grade or better. Reagent water (18 mX cm) was
D.R. Killelea, J.H. Aldstadt, III / Chemosphere 48 (2002) 1003–1008
Both core samples and grab samples (using a clam-shell device or ‘‘PONAR’’) were collected from this station. Using a boxcore sampler, a 24-cm core sample (7.64 cm diameter) was obtained and fractionated immediately in the onboard wet lab in the dark in a N2 -aspirated glove bag (4-cm fractions). The six core fractions (unacidified, unfiltered) were placed into 0.5-l opaque HDPE bottles and then immediately stored at 4 °C. The samples were stored for 180 days at 4 °C. In the laboratory, the sediment samples initially were tumbled for 24 h at 60 rpm. In a N2 atmosphere, the HDPE bottle cap was then punctured and the SPME fiber (Carboxen-PDMS (85 lm), Supelco) was equilibrated in the headspace for 60 min at 25 °C.
3. Results and discussion The herbicide DMAA was manufactured by Ansul Corporation at a plant located in Marinette, WI between 1952 and 1972 (Moyerman and Ehman, 1965). Improper disposal of industrial wastes resulted in arsenic contamination of the area, with levels in groundwater beneath a former disposal site exceeding 8500 ppm total arsenic. The chemical speciation of arsenic was studied in this region by several groups (Holm et al., 1979; Christensen and Chien, 1981). Chemical speciation was defined at that time by the determination of four major arsenic species (arsenious acid, arsenic acid, MMAA, and DMAA) near the river mouth (Holm et al., 1979) and by measurement of total arsenic in the open waters of central Green Bay (Christensen and Chien, 1981). New analytical techniques have since become available that permit a more comprehensive speciation of ‘‘arsenic’’. We therefore chose to apply our recent method to the Green Bay sediment to determine if minor arsenicals were also present. The key to studying arsines in the environment lies in the sampling stage of analysis. In general, oxidation reactions, photochemical reactions, thermal decomposition reactions, microbial changes, etc. have been well-documented as processes that can alter a sample upon collection and storage (Keith, 1996). In particular, arsines are sensitive to thermal decomposition and reaction with atmospheric oxygen (Clark, 1989) and photochemical redox reactions have also been observed for organoarsenicals (Cullen and Reimer, 1989). We therefore carefully conducted the core fractionation in the absence of light and oxygen, and then immediately stored the samples under an inert atmosphere at low temperature. The study of sediments for specific arsenic compounds has also been hindered because the chemical extractions employed often cause a loss of speciation information (Cullen and Reimer, 1989). We therefore passively sampled the headspace by SPME so as to not introduce contamination and/or artifacts that could
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arise through intensive sample manipulation. Furthermore, in dealing with the measurement of microcontaminants in complex matrices, the enrichment and selectivity capabilities that SPME headspace sampling provides were advantageous as well. The 24-cm sediment core sample was fractionated into six 4-cm segments (125 g per fraction); the headspace over each was sampled (n ¼ 4) using an 85 lm Carboxen-PDMS fiber for 60 min at room temperature. From analysis of the GC-MS response, we identified dimethylchloroarsine (DMCA) in the instrument response (Fig. 1) for a SPME headspace extract at a retention time of 3.29 min (oven temperature at this moment was 45 °C) in the core and grab samples. We base our proposed structure for this molecule on the high correlation of the m=z values (Table 1) for the two spectra (Fig. 2), which shows essentially no variation for the 15 major species. Furthermore, the relative abundances for all 22 species in the library spectrum averaged 3% of those observed for the unknown spectrum. These minute variations perhaps arise from the difference in mass analyzer used to measure the library spectrum (linear quadrupole) and the unknown spectrum (quadrupole ion trap). Unlike our previous work, we observed very little evidence of self-chemical ionization processes in the ion trap; the only species (1% abundant) identified was M þ 89 at m=z 229. The mass spectra observed across the zone width were invariant with that shown in Fig. 2(a) thereby indicating that coeluting compounds were not present. We also observed a single sharp peak at this retention time in the responses of both an electron capture detector (ECD) and a pulsed flame photometric detector (PFPD). The ECD response provides evidence for the presence of chlorine in the molecular structure (results not shown). In a similar light, the 35 Cl:37 Cl ratio in the mass spectrum (i.e., for m=z 140:142 and m=z 125:127)
Fig. 1. Portion of GC-MS chromatogram obtained for the SPME headspace extract from the Station 40 Green Bay sediment samples. The DMCA elution time on the GC-PFPD (not shown) was 30 s earlier. For conditions, see text.
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Table 1 Interpretation of the DMCA mass spectrum observed for sample species (GB) with peak area 5% that of the base peak (BP) m=z
% BPGB
% BPLIB
Process
Probable ion
75 76 88 89 90 101 102 103 105 110 124 125 127 140 142
18 5 28 85 11 17 7 31 31 11 7 100 30 84 27
13 4 29 100 15 17 7 26 28 15 4 74 26 56 19
M-65 M-64 M-52 M-51 M-50 M-39 M-38 M-37 M-35 M-30 M-16 M-15 Mi -15 M Mi
[As]þ [H–As]þ [HC–As]þ [H2 C–As]þ [H3 C–As]þ [H2 C–As–C]þ [H3 C–As–C]þ [H3 C–As–CH]þ [H3 C–As–CH3 ]þ [As–35 Cl]þ [H2 C–As–35 Cl]þ [H3 C–As–35 Cl]þ [H3 C–As–37 Cl]þ [ðCH3 Þ2 –As–35 Cl]þ [ðCH3 Þ2 –As–37 Cl]þ
Abundance values for the library spectrum (LIB) are also listed. The fragmentation process includes the subscript ‘i’ on the molecular ion (M) to denote the 37 Cl isotope.
Fig. 2. Comparison of mass spectra for (a) the unknown compound eluting at 3.28 min and (b) library spectrum of DMCA. For conditions, see text.
was 2:90 0:07, thereby indicating a single chlorine atom in the molecular structure. We also applied our recently reported method (Killelea and Aldstadt, 2001) based on the pulsed flame photometric detector (Cheskis et al., 1993) for arsenic-specific detection with high sensitivity. The method was optimized for selectivity against S-, O-, and C-containing compounds. We observed a single peak at the same retention time in the GC-PFPD chromatogram for the Station 40 headspace sample matrix (results not shown), thus indicating the presence of arsenic in the molecular structure. The quantitative aspect of our measurement is much less certain than the qualitative component. It was not possible to create a calibration model for DMCA in the
absence of a commercially available standard material. Furthermore, such a limitation also forbade determination of DMCA’s partition coefficient for the carboxen-PDMS phase coated onto the SPME fiber, i.e., the extent of pre-concentration is not known. We nevertheless semi-quantitatively estimate the concentration of DMCA using a GC-MS calibration model prepared for DMAA (i.e., as its propanethiol derivative). The PDT reduces DMAA to its arsine before cyclization, and it elutes at 35 °C (Killelea and Aldstadt, 2001). Commercially available arsines, arsine and triphenylarsine, were not chosen for the semi-quantitative procedure because they have drastically different gas-phase properties. Using the calibration model for DMAA-PDT, however, makes two key assumptions: that it has similar SPME partitioning behavior and a roughly equivalent mass response. The calibration model is based on a least-squares regression analysis: yðcountsÞ ¼ 2:46 102 ð4:61Þ ðlg=lÞ þ 6:71 103 (9:64 102 ), r2 ¼ 0:999 over the range 1–1000 lg/l (n ¼ 3 at each level). The results of applying this model are detailed in Table 2. As shown in Table 2, we observed a bimodal distribution for DMCA as a function of depth in the core sample. We speculate that this curious distribution arose from the dredging of sediment that occurred on a large scale near the former Ansul plant in Marinette in the 1990s, with the contaminated sediment dumped several miles off-shore in Green Bay (EPA, 2000). Using a deposition rate for this station estimated by 210 Pb dating at 0.47 cm/yr (Edgington, 2000), the two uppermost layers (0–8 cm) where DMCA was found correspond to the years 1982–1999. DMCA in the lower layers (17–24 cm) corresponds to deposition (1948–1964) that would have coincided with the first decade of operation of the DMAA factory. It is clear that sampling and analytical methods used in previous studies of arsenicals in Green Bay imposed limitations that made identification of only simple methylated arsenic acids possible. It is not clear how DMCA, which was studied as a chemical weapon during the First World War (Prentiss and Fisher, 1937), could be of anthropogenic (e.g., as a byproduct of the manufacture of DMAA) or biogeochemical origin (McBride and Wolfe, 1971). Methyl chloride was a starting
Table 2 Semi-quantitative estimate of DMCA concentrations in core fraction based on calibration model for DMAA-PDT Depth (cm)
Age
(DMCA) (ppb)
0–4 5–8 9–12 13–16 17–20 21–24
1990–1999 1982–1989 1973–1981 1965–1972 1956–1964 1948–1955
14.8 369 <1.7 <1.7 3.24 272
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material in the DMAA synthetic scheme at the former herbicide factory (Moyerman and Ehman, 1965), and side reactions that could produce DMCA were possible from a chemical reactivity standpoint. Furthermore, because DMCA was not found in several fractions of the sediment core that were treated in an identical fashion, generation of DMCA as an analytical artifact appears unlikely; the absence of chlorine in the analytical system further makes artifact formation improbable. The presence of DMCA 13:5 km from the apparent industrial point source is surprising given the relative instabilities of similar compounds under ambient conditions. There is little in the chemical literature to shed light on the stability of DMCA in the environment; of the chlorinated organoarsenicals that have been reported in the literature, the so-called ‘‘Lewisites’’ have been studied most extensively. ‘‘Lewisite-I’’, dichloro(2chlorvinyl)arsine, was developed as a chemical weapon by various military establishments this past century. Lewisite-I is notoriously unstable in the environment, especially toward hydrolysis (Waters and Williams, 1950); it also undergoes oxidation and thermal decomposition quite readily (Clark, 1989). For another perspective on DMCA’s reactivity, we examined the literature for information on the reactivities of (non-chlorinated) organoarsines. There appears to be widespread agreement that primary (RAsH2 ) and secondary (R2 AsH) arsines are unstable in the environment, particularly towards oxidation. However, our review of the literature on this subject suggests that the reactivity of arsines in general and in the environment in particular is poorly understood. Dimethylarsine is reported to react with molecular oxygen as follows (Raiziss and Gavron, 1923): 2ðCH3 Þ2 AsH þ 9O2 As2 O3 þ 4CO2 þ 7H2 O The oxidation of primary (and presumably secondary) arsines also produces arsenoso (–As@O) and arseno (–As@As–) compounds (Doak and Freeman, 1970). These workers also observed that the As–C bond in aliphatic arsonic acids is readily broken only by boiling with mineral acids; these workers found that action of nitric acid on MMA yielded arsenic pentoxide, formic acid, and nitrogen dioxide. Raiziss and Gavron, in contrast, reported that reaction of MMA with nitric acid yielded methylarsonic, formic, and arsenic acids, and that DMA reacted with nitric acid to form DMAA as follows: ðCH3 Þ2 AsH þ 4HNO3 ðCH3 Þ2 AsOOH þ 4NO2 þ 2H2 O
Thus, there are discrepancies in the few studies appearing in the literature related to organoarsine reactivity. We are presently attempting to better understand the distribution and abundance of DMCA in Green Bay by performing more aggressive sample pre-treatment
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of the solid phase of these samples and by comprehensively sampling sediments in the area. Additionally, we are conducting laboratory studies on the kinetics of DMCA degradation and developing an in situ monitor for organoarsines based on our previous design for chlorinated organoarsenicals (Aldstadt, 1997) to further support our studies. We have shown that headspace SPME GC-MS without thiol derivatization provides a capability for structure elucidation of organoarsines such as DMCA in complex sample matrices. The combination of careful sampling, efficient sample pre-treatment, and selective detection allowed us to identify a heretofore unobserved toxic organoarsine in this watershed.
Acknowledgements This work was supported in part by the UWM Graduate School. Presented in part at the Great Lakes Regional Meeting of the American Chemical Society (June 2000, Fargo, ND) and at the 28th Federation of Analytical Chemistry and Spectroscopy Societies Meeting (October 2001, Detroit, MI).
References Aldstadt, J.H., 1997. A flow injection trace gas analysis method for on-site determination of organoarsenicals. US Patent and Trademark Office #5,641,686, issued June 24, 1997. Anderson, L.C.D., Bruland, K.W., 1991. Biogeochemistry of arsenic in natural waters: the importance of methylated species. Environ. Sci. Technol. 25, 420–427. Andreae, M.O., 1977. Determination of arsenic species in natural waters. Anal. Chem. 49, 820–823. Andreae, M.O., 1979. Arsenic speciation in seawater and interstitial waters: the influence of biological–chemical interactions on the chemistry of a trace element. Limnol. Oceanogr. 24, 440–452. Andreae, M.O., 1986. Organoarsenic Compounds. In: Craig, P.J. (Ed.), Organometallic Compounds in the Environment. Longman Group Ltd., Essex, England, Chapter 5. Challenger, D.P., Higginbotham, C., Ellis, L., 1933. Formation of organometalloidal components by microorganisms. J. Chem. Soc., 95–101. Cheskis, S., Atar, E., Amirav, A., 1993. Pulsed-flame photomer: A novel gas Chromatography. Anal. Chem. 65, 539– 555. Christensen, E.R., Chien, N.K., 1981. Fluxes of arsenic, lead, zinc, and cadmium to Green Bay and Lake Michigan sediments. Environ. Sci. Technol. 15, 553–558. Clark, D.N., 1989. Review of Reactions of Chemical Agents in Water. Battelle Memorial Institute, Columbus, OH. Cullen, W.R., Reimer, K.J., 1989. Arsenic speciation in the environment. Chem. Rev. 89, 713–764. Doak, G.O., Freeman, L.D., 1970. In: Organometallic Compounds of Arsenic, Antimony, and Bismuth. John Wiley & Sons, New York, p. 126.
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Edgington, 2000. D.N. Personal communication. Ferguson, J.F., Gavis, J., 1972. A review of the arsenic cycle in natural waters. Water Res. 6, 1259–1274. Holm, T.R., Anderson, M.A., Iverson, D.G., Stanforth, R.S., 1979. Heterogeneous interactions of arsenic in aquatic systems. In: Chemical Modeling in Aqueous Systems. American Chemical Society, Washington, DC, pp. 711–736, Chapter 31. Environmental Protection Agency, 2000. Great Lakes National Program Office (http://www.epa.gov/grtlakes/aoc/lowmeno/ index.htm). Irgolic, K.J., 1992. The determination of arsenic and arsenic compounds in environmental samples excluding human samples. In: Stoeppler, M. (Ed.), Hazardous Metals in the Environment (Techniques and Instrumentation in Analytical Chemistry). Elsevier, Amsterdam, Chapter 11. Keith, L.H., 1996. Principles of Environmental Sampling, 2nd ed. American Chemical Society, Washington, DC. Killelea, D.R., Aldstadt, J.H., 2001. A solid-phase microextraction method for gas chromatography with mass spectrometric and pulsed flame photometric detection: studies of organoarsenical speciation. J. Chromatogr. 918A, 169– 175. Le, X.C., Ma, M., 1998. Short-column liquid chromatography with hydride generation atomic fluorescence detection for the speciation of arsenic. Anal. Chem. 70, 1926–1933.
Matschullat, J., 2000. Arsenic in the geosphere––a review. Sci. Total Environ. 249, 297–312. McBride, B.C., Wolfe, R.S., 1971. Biosynthesis of dimethylarsine by methanobacterium. Biochemistry 10, 4317–4321. Moyerman, R.M., Ehman, P.J., 1965. Manufacture of arsinic acids. US Patent and Trademark Office #3,173,937, issued March 6, 1965. Pantsar-Kallio, M., Korpela, A., 2000. Analysis of gaseous arsenic species and stability studies of arsine and trimethylarsine by gas chromatograpy-mass spectrometry. Anal. Chim. Acta 410, 65–70. Pawliszyn, J., 1997. Solid Phase Microextraction: Theory and Practice. Wiley-VCH, New York. Penrose, W.R., 1974. Arsenic in the marine and aquatic environments: analysis, occurrence, and significance. CRC Crit. Rev. Environ. Control, 465–482. Prentiss, A.M., Fisher, G.J.B., 1937. Chemicals in War: A Treatise on Chemical Warfare. McGraw-Hill, New York. Raiziss, G.W., Gavron, J.L., 1923. In: Organic Arsenical Compounds. The Chemical Catalog Company, p. 38. Szostek, B., Aldstadt, J.H., 1998. Determination of organoarsenicals in the environment by solid-phase microextraction gas chromatography-mass spectrometry. J. Chromatogr. 807A, 253–263. Waters, W.A., Williams, J.H., 1950. Hydrolyses and derivatives of some vesicant arsenicals. J. Chem. Soc., 18–22.
Identification of dimethylchloroarsine near a former herbicide factory by headspace solid-phase microextraction gas chromatography-mass spectrometry Daniel R. Killelea 1, Joseph H. Aldstadt, III
*
Department of Chemistry, University of Wisconsin-Milwaukee, 3210 North Cramer Street, Milwaukee, WI 53211, USA Received 7 December 2000; received in revised form 27 March 2002; accepted 30 March 2002
Abstract The application of an improved method based on multidetector gas chromatography to the determination of trace levels of organoarsines in complex matrices is described. The method using headspace-mode solid-phase microextraction (SPME) was applied to a carefully sampled and preserved freshwater sediment core obtained from central Green Bay, Lake Michigan. The sediment core was collected and fractionated in an inert atmosphere. A carboxen/ polydimethylsiloxane-coated SPME fiber (85 lm film thickness) was equilibrated (n ¼ 4) for 60 min at 25 °C in the headspace of the sample vessel before introduction to the chromatograph. Conventional quadrupole ion trap mass spectrometry (electron impact ionization), electron capture detection, and pulsed flame photometric detection (arsenic mode) were employed for structure elucidation. A heretofore unidentified species in this region, dimethylchloroarsine (DMCA), was identified. The mass spectrum for DMCA is interpreted based on the observed fragmentation pattern. A bimodal vertical distribution of DMCA in the sediment core sample was observed and its interpretation based on Pb210 dating is reported. Ó 2002 Published by Elsevier Science Ltd. Keywords: Arsenic; Green bay; Dimethylchloroarsine [CAS registry no. 557-89-1]
1. Introduction Arsenic is ubiquitous in nature, and it is now firmly established that the impact of environmental arsenic cannot be expressed simply as a function of total arsenic concentration (Ferguson and Gavis, 1972). Indeed, the many compounds that this reactive metalloid can form encompass a wide range of physicochemical and toxicological properties (Matschullat, 2000). To better understand transport and ultimate effect on a given ecosystem, it is critical to identify and quantify all of the specific
*
Corresponding authors. Tel.: +414-229-5605; fax: +414229-5530. E-mail address: [email protected] (J.H. Aldstadt, III). 1 Present address: Department of Chemistry, Tufts University, 62 Talbot Avenue, Medford, MA 02155, USA.
chemical forms of arsenic that are present (Cullen and Reimer, 1989). Providing accurate information on the abundance, distribution, and structures of arsenicals at environmentally relevant levels is analytically challenging. The chemical species of arsenic in the environment are typically divided into five general classes: (a) inorganic covalent oxygen and sulfur forms, such as As2 S3 and As2 O3 ; (b) the inorganic protonated oxyanions, i.e., arsenious acid, As(OH)3 , and arsenic acid, AsO(OH)3 ; (c) alkylated derivatives of the acids, including, monomethylarsonic acid (MMAA), dimethylarsinic acid (DMAA), trimethyl arsine oxide, arsenobetaine, and arsenocholine; (d) the so-called ‘‘arsenosugars’’, with most attention focusing on the major species found in marine organisms; and (e) the arsines (Penrose, 1974; Andreae, 1977; Andreae, 1979; Anderson and Bruland, 1991).
0045-6535/02/$ - see front matter Ó 2002 Published by Elsevier Science Ltd. PII: S 0 0 4 5 - 6 5 3 5 ( 0 2 ) 0 0 1 6 0 - 1
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The major forms of arsenic that fall into the ‘‘arsine’’ category are inorganic arsine (AsH3 ) and aliphatic (methylated) derivatives of trivalent arsenic––monomethyl arsine (MMA), dimethyl arsine (DMA), and trimethyl arsine). Arsines can be generated by bacterial or fungal action, e.g., the classic example is the action of the fungus Scopulariopsis brevicaulis in producing trimethylarsine (Challenger et al., 1933; Andreae, 1986). As a class, the arsines represent some of the most toxic forms of arsenic, and under certain conditions can be easily transported in nature because of their pronounced volatility. At the same time, the arsines are generally reactive toward hydrolysis and oxidation, thus in many instances mitigating their potentially detrimental effects. While methods for measuring total arsenic and most major arsenic species are numerous (Irgolic, 1992), there are relatively few modern methods for analysis of the arsines in environmental samples (Le and Ma, 1998; Pantsar-Kallio and Korpela, 2000). Equally importantly, there is substantial inconsistency in the procedures used to collect arsenicals in the field in general and arsines in particular prior to their determination in the laboratory (Cullen and Reimer, 1989). The difficulty in measuring the arsines usually arises in the sampling phase of analysis because one must exercise great care to accurately sample such volatile and potentially reactive compounds. Recently we described a method for the determination of several organoarsenic acids that combined chemical derivatization with solid-phase microextraction (SPME) and gas chromatography-mass spectrometry (GC-MS) (Szostek and Aldstadt, 1998; Killelea and Aldstadt, 2001). For polar and/or thermallylabile analytes, analyte derivatization prior to SPME-GC can enhance detectability and improve chromatographic resolution (Pawliszyn, 1997). We examined a series of dithiol compounds for derivatization of the arsenicals, and the best results were obtained with 1,3-propanedithiol (PDT). The derivatization procedure, fiber type, and extraction time were then optimized. Using this method, the limit of detection for 2-chlorovinyl arsonous acid was improved 400 fold compared to conventional solvent extraction methods. The goal of the work reported herein was to modify our previous SPME GC-MS method (Killelea and Aldstadt, 1998) by using headspace-mode SPME without thiol derivatization to carefully preserved freshwater sediments near a former herbicide manufacturing facility to determine if organoarsines were present.
prepared using a NanoPureTM filtration system equipped with an ultraviolet lamp (Barnstead-Thermolyne, Dubuque, IA, USA). Arsenic standards were obtained from Aldrich Chemical (Milwaukee, WI, USA). All glassware and plasticware were acid-washed after use for at least 48 h in 5% (v/v) AR-grade nitric acid to remove background contamination. Arsenic-containing standards with concentrations less than 1 mg/l were prepared on the day of use. All aqueous standards were stored at 4 °C. 2.2. Instrumentation Organoarsenicals were determined by gas–liquid chromatography using a Varian GC-MS system (Walnut Creek, CA, USA) located in a Class 100 clean room. The GC-MS consisted of the following components: Model 3800 capillary gas–liquid chromatograph (DB5MS column, 30 m 0:25 mm with 0.25 lm film, J&W Scientific, Folsom, CA, USA) with Model 1079 split/ splitless injector; SPME apparatus (Supelco, Bellefonte, PA, USA); electron impact ionization source (70 eV); Saturn 2000 quadrupole ion trap MS (10–650 m=z range, unit resolution). The automatic gain control of the MS system was used throughout this study. Automated library searching was performed using the NIST Mass Spectral Database (version 3.0). The mobile phase was ultrahigh purity (99.999%) helium (Praxair, Milwaukee, WI, USA) at a constant linear velocity of 42 cm/s by electronic flow control. Samples were also studied using electron capture detection and pulsed flame photometric detection, as described elsewhere (Killelea and Aldstadt, 2001). 2.3. GC program For the SPME work, the injection port was 250 °C for the fiber desorption step (splitless). The initial column temperature was 45 °C for 5 min, then programmed at 20 °C/min to 160 °C, followed by 8 °C/min to 210 °C and finally 50 °C/min to 300 °C, where it was held for 5 min. The transfer line between the GC and MS was maintained at 170°. Mass spectra were obtained by scanning from m=z 35 to 400 with a 0.7 s scan time. GC instrument control and data acquisition were performed on a Pentium personal computer (Dell, Optiplex GX1, Dallas, TX, USA) using Saturn System software version 5.21 (Varian).
2. Experimental
2.4. Sediment procedures
2.1. Reagents
Sediment samples were collected in July 1999 aboard R/V Neeskay at a depth of approximately 20 m in central Green Bay, approximately 13.5 km south of the mouth of the Menominee River (Green Bay Station 40).
All chemical reagents used were analytical reagent (AR) grade or better. Reagent water (18 mX cm) was
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Both core samples and grab samples (using a clam-shell device or ‘‘PONAR’’) were collected from this station. Using a boxcore sampler, a 24-cm core sample (7.64 cm diameter) was obtained and fractionated immediately in the onboard wet lab in the dark in a N2 -aspirated glove bag (4-cm fractions). The six core fractions (unacidified, unfiltered) were placed into 0.5-l opaque HDPE bottles and then immediately stored at 4 °C. The samples were stored for 180 days at 4 °C. In the laboratory, the sediment samples initially were tumbled for 24 h at 60 rpm. In a N2 atmosphere, the HDPE bottle cap was then punctured and the SPME fiber (Carboxen-PDMS (85 lm), Supelco) was equilibrated in the headspace for 60 min at 25 °C.
3. Results and discussion The herbicide DMAA was manufactured by Ansul Corporation at a plant located in Marinette, WI between 1952 and 1972 (Moyerman and Ehman, 1965). Improper disposal of industrial wastes resulted in arsenic contamination of the area, with levels in groundwater beneath a former disposal site exceeding 8500 ppm total arsenic. The chemical speciation of arsenic was studied in this region by several groups (Holm et al., 1979; Christensen and Chien, 1981). Chemical speciation was defined at that time by the determination of four major arsenic species (arsenious acid, arsenic acid, MMAA, and DMAA) near the river mouth (Holm et al., 1979) and by measurement of total arsenic in the open waters of central Green Bay (Christensen and Chien, 1981). New analytical techniques have since become available that permit a more comprehensive speciation of ‘‘arsenic’’. We therefore chose to apply our recent method to the Green Bay sediment to determine if minor arsenicals were also present. The key to studying arsines in the environment lies in the sampling stage of analysis. In general, oxidation reactions, photochemical reactions, thermal decomposition reactions, microbial changes, etc. have been well-documented as processes that can alter a sample upon collection and storage (Keith, 1996). In particular, arsines are sensitive to thermal decomposition and reaction with atmospheric oxygen (Clark, 1989) and photochemical redox reactions have also been observed for organoarsenicals (Cullen and Reimer, 1989). We therefore carefully conducted the core fractionation in the absence of light and oxygen, and then immediately stored the samples under an inert atmosphere at low temperature. The study of sediments for specific arsenic compounds has also been hindered because the chemical extractions employed often cause a loss of speciation information (Cullen and Reimer, 1989). We therefore passively sampled the headspace by SPME so as to not introduce contamination and/or artifacts that could
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arise through intensive sample manipulation. Furthermore, in dealing with the measurement of microcontaminants in complex matrices, the enrichment and selectivity capabilities that SPME headspace sampling provides were advantageous as well. The 24-cm sediment core sample was fractionated into six 4-cm segments (125 g per fraction); the headspace over each was sampled (n ¼ 4) using an 85 lm Carboxen-PDMS fiber for 60 min at room temperature. From analysis of the GC-MS response, we identified dimethylchloroarsine (DMCA) in the instrument response (Fig. 1) for a SPME headspace extract at a retention time of 3.29 min (oven temperature at this moment was 45 °C) in the core and grab samples. We base our proposed structure for this molecule on the high correlation of the m=z values (Table 1) for the two spectra (Fig. 2), which shows essentially no variation for the 15 major species. Furthermore, the relative abundances for all 22 species in the library spectrum averaged 3% of those observed for the unknown spectrum. These minute variations perhaps arise from the difference in mass analyzer used to measure the library spectrum (linear quadrupole) and the unknown spectrum (quadrupole ion trap). Unlike our previous work, we observed very little evidence of self-chemical ionization processes in the ion trap; the only species (1% abundant) identified was M þ 89 at m=z 229. The mass spectra observed across the zone width were invariant with that shown in Fig. 2(a) thereby indicating that coeluting compounds were not present. We also observed a single sharp peak at this retention time in the responses of both an electron capture detector (ECD) and a pulsed flame photometric detector (PFPD). The ECD response provides evidence for the presence of chlorine in the molecular structure (results not shown). In a similar light, the 35 Cl:37 Cl ratio in the mass spectrum (i.e., for m=z 140:142 and m=z 125:127)
Fig. 1. Portion of GC-MS chromatogram obtained for the SPME headspace extract from the Station 40 Green Bay sediment samples. The DMCA elution time on the GC-PFPD (not shown) was 30 s earlier. For conditions, see text.
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Table 1 Interpretation of the DMCA mass spectrum observed for sample species (GB) with peak area 5% that of the base peak (BP) m=z
% BPGB
% BPLIB
Process
Probable ion
75 76 88 89 90 101 102 103 105 110 124 125 127 140 142
18 5 28 85 11 17 7 31 31 11 7 100 30 84 27
13 4 29 100 15 17 7 26 28 15 4 74 26 56 19
M-65 M-64 M-52 M-51 M-50 M-39 M-38 M-37 M-35 M-30 M-16 M-15 Mi -15 M Mi
[As]þ [H–As]þ [HC–As]þ [H2 C–As]þ [H3 C–As]þ [H2 C–As–C]þ [H3 C–As–C]þ [H3 C–As–CH]þ [H3 C–As–CH3 ]þ [As–35 Cl]þ [H2 C–As–35 Cl]þ [H3 C–As–35 Cl]þ [H3 C–As–37 Cl]þ [ðCH3 Þ2 –As–35 Cl]þ [ðCH3 Þ2 –As–37 Cl]þ
Abundance values for the library spectrum (LIB) are also listed. The fragmentation process includes the subscript ‘i’ on the molecular ion (M) to denote the 37 Cl isotope.
Fig. 2. Comparison of mass spectra for (a) the unknown compound eluting at 3.28 min and (b) library spectrum of DMCA. For conditions, see text.
was 2:90 0:07, thereby indicating a single chlorine atom in the molecular structure. We also applied our recently reported method (Killelea and Aldstadt, 2001) based on the pulsed flame photometric detector (Cheskis et al., 1993) for arsenic-specific detection with high sensitivity. The method was optimized for selectivity against S-, O-, and C-containing compounds. We observed a single peak at the same retention time in the GC-PFPD chromatogram for the Station 40 headspace sample matrix (results not shown), thus indicating the presence of arsenic in the molecular structure. The quantitative aspect of our measurement is much less certain than the qualitative component. It was not possible to create a calibration model for DMCA in the
absence of a commercially available standard material. Furthermore, such a limitation also forbade determination of DMCA’s partition coefficient for the carboxen-PDMS phase coated onto the SPME fiber, i.e., the extent of pre-concentration is not known. We nevertheless semi-quantitatively estimate the concentration of DMCA using a GC-MS calibration model prepared for DMAA (i.e., as its propanethiol derivative). The PDT reduces DMAA to its arsine before cyclization, and it elutes at 35 °C (Killelea and Aldstadt, 2001). Commercially available arsines, arsine and triphenylarsine, were not chosen for the semi-quantitative procedure because they have drastically different gas-phase properties. Using the calibration model for DMAA-PDT, however, makes two key assumptions: that it has similar SPME partitioning behavior and a roughly equivalent mass response. The calibration model is based on a least-squares regression analysis: yðcountsÞ ¼ 2:46 102 ð4:61Þ ðlg=lÞ þ 6:71 103 (9:64 102 ), r2 ¼ 0:999 over the range 1–1000 lg/l (n ¼ 3 at each level). The results of applying this model are detailed in Table 2. As shown in Table 2, we observed a bimodal distribution for DMCA as a function of depth in the core sample. We speculate that this curious distribution arose from the dredging of sediment that occurred on a large scale near the former Ansul plant in Marinette in the 1990s, with the contaminated sediment dumped several miles off-shore in Green Bay (EPA, 2000). Using a deposition rate for this station estimated by 210 Pb dating at 0.47 cm/yr (Edgington, 2000), the two uppermost layers (0–8 cm) where DMCA was found correspond to the years 1982–1999. DMCA in the lower layers (17–24 cm) corresponds to deposition (1948–1964) that would have coincided with the first decade of operation of the DMAA factory. It is clear that sampling and analytical methods used in previous studies of arsenicals in Green Bay imposed limitations that made identification of only simple methylated arsenic acids possible. It is not clear how DMCA, which was studied as a chemical weapon during the First World War (Prentiss and Fisher, 1937), could be of anthropogenic (e.g., as a byproduct of the manufacture of DMAA) or biogeochemical origin (McBride and Wolfe, 1971). Methyl chloride was a starting
Table 2 Semi-quantitative estimate of DMCA concentrations in core fraction based on calibration model for DMAA-PDT Depth (cm)
Age
(DMCA) (ppb)
0–4 5–8 9–12 13–16 17–20 21–24
1990–1999 1982–1989 1973–1981 1965–1972 1956–1964 1948–1955
14.8 369 <1.7 <1.7 3.24 272
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material in the DMAA synthetic scheme at the former herbicide factory (Moyerman and Ehman, 1965), and side reactions that could produce DMCA were possible from a chemical reactivity standpoint. Furthermore, because DMCA was not found in several fractions of the sediment core that were treated in an identical fashion, generation of DMCA as an analytical artifact appears unlikely; the absence of chlorine in the analytical system further makes artifact formation improbable. The presence of DMCA 13:5 km from the apparent industrial point source is surprising given the relative instabilities of similar compounds under ambient conditions. There is little in the chemical literature to shed light on the stability of DMCA in the environment; of the chlorinated organoarsenicals that have been reported in the literature, the so-called ‘‘Lewisites’’ have been studied most extensively. ‘‘Lewisite-I’’, dichloro(2chlorvinyl)arsine, was developed as a chemical weapon by various military establishments this past century. Lewisite-I is notoriously unstable in the environment, especially toward hydrolysis (Waters and Williams, 1950); it also undergoes oxidation and thermal decomposition quite readily (Clark, 1989). For another perspective on DMCA’s reactivity, we examined the literature for information on the reactivities of (non-chlorinated) organoarsines. There appears to be widespread agreement that primary (RAsH2 ) and secondary (R2 AsH) arsines are unstable in the environment, particularly towards oxidation. However, our review of the literature on this subject suggests that the reactivity of arsines in general and in the environment in particular is poorly understood. Dimethylarsine is reported to react with molecular oxygen as follows (Raiziss and Gavron, 1923): 2ðCH3 Þ2 AsH þ 9O2 As2 O3 þ 4CO2 þ 7H2 O The oxidation of primary (and presumably secondary) arsines also produces arsenoso (–As@O) and arseno (–As@As–) compounds (Doak and Freeman, 1970). These workers also observed that the As–C bond in aliphatic arsonic acids is readily broken only by boiling with mineral acids; these workers found that action of nitric acid on MMA yielded arsenic pentoxide, formic acid, and nitrogen dioxide. Raiziss and Gavron, in contrast, reported that reaction of MMA with nitric acid yielded methylarsonic, formic, and arsenic acids, and that DMA reacted with nitric acid to form DMAA as follows: ðCH3 Þ2 AsH þ 4HNO3 ðCH3 Þ2 AsOOH þ 4NO2 þ 2H2 O
Thus, there are discrepancies in the few studies appearing in the literature related to organoarsine reactivity. We are presently attempting to better understand the distribution and abundance of DMCA in Green Bay by performing more aggressive sample pre-treatment
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of the solid phase of these samples and by comprehensively sampling sediments in the area. Additionally, we are conducting laboratory studies on the kinetics of DMCA degradation and developing an in situ monitor for organoarsines based on our previous design for chlorinated organoarsenicals (Aldstadt, 1997) to further support our studies. We have shown that headspace SPME GC-MS without thiol derivatization provides a capability for structure elucidation of organoarsines such as DMCA in complex sample matrices. The combination of careful sampling, efficient sample pre-treatment, and selective detection allowed us to identify a heretofore unobserved toxic organoarsine in this watershed.
Acknowledgements This work was supported in part by the UWM Graduate School. Presented in part at the Great Lakes Regional Meeting of the American Chemical Society (June 2000, Fargo, ND) and at the 28th Federation of Analytical Chemistry and Spectroscopy Societies Meeting (October 2001, Detroit, MI).
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