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LITERATURE CITED Ettore Santoro for suggestions and helpful discussions. Leonhardt, 8. A.; De Viibiss, E. D.; Klun, J. A. Org. Mass Spectrom. Registry No. Z,Z-7,13-C18Ac,105835-82-3;Z,Z-3,9-C18Ac, 1983, 78, 9-11. 105835-83-4; E,Z-2,9-C18Ac, 105835-84-5; Z,Z-2,13-C18A~, Lanne, B. S.;Appelgren, M.; Bergstrom, G.; Lofstedt, C. Anal. Chem. 86252-65-5; E,Z-2,13-C18Ac, 86252-74-6; Z,E-2,13-C18A~, 1985, 57, 1621-1625. Hogge, L. R.; Olson, D. J. H. J. Chromafcgr. Sci. 1983, 27,524-528. 102637-06-9; E,E-2,13-C18Ac, 105835-85-6; Z,Z-3,13-C18Ac, Peake. D. A.; Gross, M. L. Anal. Chem. 1985, 57, 115-120. 53120-27-7; Z,Z-3,14-C18Ac, 86252-66-6; Z,Z-3,12-C18Ac, Jensen, N. J. J.; Tomer, K. B.; Gross, M. L. Anal. Chem. 1985, 57, 105835-86-7; E,Z-3,13-C18Alc, 66410-28-4; Z,Z-5,11-C16Al~, 2016-2021. 105835-87-8; Z,Z-5,9-C18Ac, 105835-88-9; Z,Z-6,9-C18Ac, Ferrer-Correia, A. J. V.; Jennings, K. R.; Sen Sharma, D. K. Org. Mass Specfrom. 1976, 7 7 , 867-872. 105835-89-0; Z,Z-4,9-C18Ac, 105835-90-3; Z,Z-3,8-C18Ac, Chai, R.; Harrison, A. G. Anal. Chem. 1981, 53,34-37. 105835-91-4; Z,Z-7,11-C16A~,52207-99-5; Z,E-7,11-C16Ac, Ghaderi, S.;Kuikarni, P. S.; Ledford, E. B.; Wilkins, C. L.; Gross, M. L. 53042-79-8; Z,Z-5,9-C18Alc, 105835-92-5; Z,Z-8,12-C18Est, Anal. Chem. 1981, 53,428-437. Budzikiewicz, H.; Busker, E. Tetrahedron 1980, 36, 255-266. 26258-13-9;Z,Z7,9-C18Ac,105835-93-6;E,Z3,5-€14Ac, 61360-85-8; Doolittle, R. E.: Tumlinson, J. H.; Proveaux, A. Anal. Chem. 1985, 57, E-9,11-C12Ac, 50767-78-7; 1,7-octadiene, 3710-30-3; dimethyl 1625- 1630. disulfide,62492-0; 7,8,13,14tetramethylthi~decan-l-ol acetate, Brauner. A.; Budzikiewlcz, H.; Boland, W. Org . Mass Spectrom . 1982, 105835-94-7; 3,4,9,10-tetramethylthiooctadecan-l-olacetate, 77,161-164. Bambagiotti, M. A.; Coran, S. A.; Giannellini, V.; Vincieri, F. F.; Daoiio, 105835-95-8; 2,3,9,10-tetramethylthiooctadecan-l-olacetate, S.;Traldi, P. Org. hhss Specfrom. W83, 78, 133-134. 105835-96-9; 2,3,13,14-tetramethylthiooctadecan-l-ol acetate, Bambagiotti, M. A.; Coran, S. A,; Gianneliini, V.; Vincieri, F. F.; Daolio, 105835-97-0; 3,4,13,14-tetramethylthiooctadecan-l-ol acetate, S.;Traldi, P. Org. Chem. Spectrom. 1984, 79,577-580. 105835-98-1; 3,4,14,15-tetramethylthiooctadecan-l-olacetate, Kidweii, D. A.: Biemann, K. Anal. Chem. 1982, 54,2462-2465. Cervilla, M.; Puzo, G. Anal. Chem. 1983, 55,2100-2103. 105835-99-2; 3,4,12,13-tetramethylthiooctadecan-l-ol acetate, Francis, G. W.; Veiand, K. J. Chromatogr. 1961, 279, 379-384. 105836-00-8;5,6,11,12-tetramethylthiohexadecan-l-o1, 105836-01-9; Buser, H. R.; Arn, H.; Guerin, P.; Rauscher, S.Anal. Chem. 1983, 55, 3,4,13,14-tetramethylthiooctadecan-l-ol, 105836-02-0; 1,2,7,8818-822. tetramethylthiooctadiene, 105836-03-1;2-(1-acetoxy-5-methylDunkelblum, E.; Tan, S. H.; Silk, P. J. J . Chem. Ecol. 1985, 7 7 , 265-277. thiopentan-5-y1)-5-(1-methylthiononan-1-yl)tetrahydrothiophene, Leonhardt, B. A.; De Vilbiss, E. D. J. Chromatogr. 1985, 322, 105836-04-2; 2-(l-acetoxy-6-methylthiohexan-6-yl)-4-(l-methyl484-490. thiononan-1-yl)thietane,105836-05-3; 2-(1-acetoxy-4-methylNiehaus, W. G.,Jr.; Ryhage, R. Anal. Chem. 1968, 4 0 , 1840-1847. thiobutan-4-~1)-6-( l-methylthiononan-l-yl)-2H-tetrahydrothioHogge, L. R.; Underhiil, E. W.; Wong, J. W. J. Chromatogr. Sci. 1985, 23, 171-175. pyran, 105836-06-4; 2-(l-acetoxy-3-methylthioprop-3-yl)-6-(l413-421. Ando, T.; Katagiri, Y.; Uchiyama, M. Agric. Biol. Chem. 1985, 49, methylthiodecan-l-yl)-2H-tetrahydrothiopyran,105836-07-5; 2-(l-acetoxy-7-methylthioheptan-7-yl)-5-(l-methylthiopentan-lCaserio, M. C.; Fischer, C. L.; Kim, J. K. J. Org. Chem. 1985, 5 0 , 4390-4393. yl)tetrahydrothiophene, 105836-08-6; 2-(1-hydroxy-5-methylTonini, C.; Cassani, G.; Massardo, P.; Guglieimetti, G.; Casteiiari, P. L. thiopentan-5-~1)-5-( 1-methylthiononan-1-yl)tetrahydrothiophene, J . Chem. Ecol. 1986, 72,1545-1556. 105836-09-7; 2-(methyl-8-rnethylthiooctanoate-8-yl)-5-(1methylthiohexan-1-yl)tetrahydrothiophene,105836-10-0;241for review May 29,1986. Accepted October 20,1986. acetoxyoctan-8-yl)-3,4-dimethylthiotetrahydrothiophene, RECEIVED 105836-11-1; 2-(l-acetoxyethan-2-yl)-3,4-dimethylthio-5-(octan- Part of this work had been presented a t the A. J. P. Martin 1-yl)tetrahydrothiophene, 105836-12-2. Honorary Symposium (Urbino, Italy, May 27-31, 1985).
Stability of Hydrocarbon Samples on Solid-Phase Extraction Columns David R. Green*l and Donna
Le Pape
Seakem Oceanography Ltd., 2045 Mills Road, Sidney, British Columbia, Canada V8L 35'1
The stabillty of hydrocarbon samples sorbed from water onto two types of solid phases was examined. The two solid phases, X A P P macroreticular resln and octadecane bonded on silica gel, were found to have a preservative effect which prevented the breakdown of sorbed hydrocarbons by bacteria. Hydrocarbons stored on these solid phases for perlods of up to 100 days In the presence of an oieophllic bacterial population showed no evldence of biological degradation as lndlcated by changes in chromatographic pattern or degradation of a radiolabeled hydrocarbon. I n contrast, hydrocarbons stored in water samples containing the same bacteria showed pronounced degradation over much shorter storage periods. The macroreticular or pore structure of the solid phases is thought to be the mechanism by which the extracted hydrocarbons are preserved from bacterial attack. Present address: Seakem Oceanography Ltd., Argo Building, B e d f o r d Institute of Oceanography, P.O. B o x 696, Dartmouth, N o v a Scotia, Canada B 2 Y 3Y9.
Determination of trace organics in water has benefited from remarkable improvements in analytical methodology over the last decade. However, sampling methodology has progressed scarcely at all and the problems identified a decade ago (nonrepresentative samples, insufficient volumes, contamination, undocumented preservation techniques, labor intensive extraction procedures) remain largely unsolved ( I ) . The use of solid-phase extraction columns to concentrate trace organics from water presents solutions to some of these problems (2, 3). The technique is less labor intensive than solvent extraction and enables concentration of organics from larger volumes of water, permitting improvements in analytical precision and accuracy. However, the real advantages of the use of solid-phase sorbents become apparent when the extraction columns are deployed in the field by using newly available submersible instrumentation ( 4 , 5 ,6). With field deployment, extraction is done in situ and the collection of the water itself is avoided, thereby eliminating most contamination and handling problems.
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The stability of samples collected on solid-phase extraction columns has not previously been studied and is of fundamental importance to the continued use of this technique of water sampling for organic analyses. The purpose of the research reported here was t o determine the fate of typical organic samples sorbed on solid phase extraction columns, with a view to developing preservation methods for column samples analogous to preservation techniques commonly used for water samples. This study investigated the stability of crude oil samples adsorbed on XAD-2 macroreticular resin and on octadecane bonded on silica gel (CIs-silica gel), two commonly used sorbents for the extraction of organics from water. Crude oil was used because it contains a wide range of organic compounds and shows well-established degradation patterns characteristic of biological degradation. The experiments were conducted under both laboratory and field conditions with oleophilic bacteria added to the columns t o promote degradation. EXPERIMENTAL SECTION Extraction Columns. XAD-2 is a polystyrene-divinylbenzene copolymer ( R o b & Haas). The resin has a mesh size of 20/60, a surface area of 300 m2/g, and a pore diameter of 90 A. XAD-2 extraction columns from a commercially available in situ sampling system (Seastar Instruments, Sidney, BC, Canada) were used in the study. The columns contained approximately 90mL of XAD-2 in a cylinder made of Teflon (2.5 cm o.d., 1.9 cm id., 37 cm long). Resin was held in place by mesh made of Teflon. Octadecane bonded to 35-70 mesh silica gel (BDH) was synthesized for the study following Sander and Wise (7). In-house synthesis allowed the selection of appropriate silica particle diameters to obtain the same column configuration and flow rates as the XAD-2 column. Prior to use both XAD-2 and C18-silica gel columns were cleaned by pumping fresh distilled methanol through them at 100 mL/min for 24 h, dichloromethane for a further 24 h, and methanol for a final rinse. Columns were stored moist with methanol and sealed with plugs made of Teflon. Prior to being loaded with hydrocarbon samples, columns were prerinsed by pumping 2 L of 95 “C distilled water through them to remove residual methanol in case the residual methanol caused a preservation effect. The crude oil used in the experiments was Arabian light crude oil (American Petroleum Institute Standard Reference Oil Program). All solvents were distilled in glass. Preparation of Standard Columns for Storage Experiments. In order to prepare columns with standard hydrocarbon and bacterial loadings for each of the storage experiments, an “input vat” containing 50 L of seawater (Instant Ocean, 20% for the first three experiments, natural seawater for the last three) was set up with a submersible pump providing mixing energy. Crude oil was added to the vat as a 1O:l oil/dispersant mixture. The dispersant (Exxon’s Corexit 9527) was necessary to stabilize the crude oil emulsion. One to two liters of a culture of marine bacteria maintained on the oil was also added to the input vat. The vat was stirred continuously for 12 h to allow equilibration of oil concentrations and adaption of the bacteria. Concentrations in the input vat for each of the six experiments a t the time of ”loading” the columns were approximately 1mg/L crude oil and 104-106 bacteria/mL. Four liters of the mixture was pumped (200 mL/min) through each of the columns. All of the columns required for an experiment could be prepared in about 2 h from one input vat. Samples of the input mixture were taken a t the beginning, middle, and end of the loading procedure and analyzed for hydrocarbon concentration and composition. No significant change in the concentration or composition of the hydrocarbons occurred over the 2-h loading periods. Alkane Composition. Bacterial degradation of oil typically results in a dramatic change in alkane composition visible on gas chromatographic traces (8,9). In particular, changes in the ratios of pristane:C,,, phytane:C18, and unresolved complex mixture (UCM):CI8 provide a measure of bacterial degradation. The
straight chain alkanes (C17and C18)are more easily degraded by bacteria than either the branched chain isoprenoids (pristane and phytane) or the UCM, and as a result, the ratios increase toward infinity during biological degradation. In order to determine the indicator ratios, the following gas chromatographic analysis of samples was conducted. Hydrocarbons on columns were eluted first with 200 mL of methanol (subsequently back extracted into three 100-mLportions of pentane) and then with 200 mL of dichloromethane. Pentane and dichloromethane fractions were combined, water was removed with granular, anhydrous sodium sulfate, and the extract was taken to near dryness. The residue was taken up in 1:1 2propanol/dichloromethane. The alkane fraction was separated from the aromatic fraction by application to a Sephadex LH20 column (Pharmacia Fine Chemicals, Uppsala, Sweden) and elution with 1:l 2-propanol/dichloromethane. The alkane fraction was taken to near dryness, applied to a 5% deactivated silica gel column, eluted with pentane, and analyzed by glass capillary gas chromatography with flame ionization detection. A HewlettPackard 5840A instrument with a DB5 capillary column was used. A perdeuteriated internal standard, n-Cs6-d7,, was carried throughout the procedure. Ratios of individual components were obtained by manual peak height measurement. Hydrocarbons in water samples (1L) were extracted with three 100-mL portions of dichloromethane. Walls of storage containers were rinsed with solvent. Combined extracts were dried over sodium sulfate, concentrated, separated, and analyzed as for the column extracts. Radioisotope Methods. Bacterial metabolism of hydrocarbons results in the generation of COP. Addition of radiolabeled hydrocarbon [ l-14C]hexadecaneto input mixtures and subsequent monitoring of the production of radiolabeled CO, and disappearance of 14C-labeled hexadecane was used as a means of measuring the rate of biological degradation (10). Concentrations of [ l-14C]hexadecanein initial and incubated water samples were determined by placing half of the organic extract used for alkane spectra determinations in Aquasol fluor (New England Nuclear) and counting on a Beckman LS 8100 scintillation counter. The 14C02was analyzed by scintillation counting after removing it from the water by placing 250-mL subsamples in a vacuum line between C02traps containing 0.5 M NaOH. Samples were acidified with 2 mL of concentrated sulfuric acid. The downstream trap collected the 14C02released from the sample, and I4CO3was precipitated out by using BaC1, and collected on two 25-mm glass-fiber filters under vacuum filtration. The filters were placed in scintillation vials with 7 mL of Aquasol, the vials were shaken until the mixture was homogeneous, and then 3 mL of water was added to form a gel to keep the precipitate in suspension. The sample was then counted. Columns were analyzed for [1-’4C]hexadecaneas above by using extracts from the alkane analysis. The 14C02in columns was determined as above from residual water contained in the column during storage and flushed out with distilled water prior to analysis. Microbiological Methods. Bacterial counts were made routinely on the input mixture, water samples, sample columns, and the bacterial culture used to supply bacteria for experiments. Bacterial populations were estimated by plating on nutrient marine agar plates (DIFCO) No. 2216) and by a microscope direct count with a Petroff-Hauser Counting Chamber. Phase contrast microscopy was used to determine if cells counted by the Petroff-Hauser technique were viable. Counts on water samples were made by collecting approximately 10 mL by sterile technique and standard dilution procedures. Counts were made the same day they were collected. For bacterial counts on sample columns, 1g of resin from the input end of the column was shaken in a vial with 10 mL of sterile water of which 1 mL was subsequently diluted and plated. Bacterial numbers were expressed as counts per unit dry weight of resin. The ability of the culture to degrade oil was also documented prior to each experiment by adding fresh oil to the culture and observing changes in the alkane composition known to be characteristic of bacterid degradation. Hydrocarbon composition was determined by methods described above. Laboratory Storage of Hydrocarbons on XAD-2 Columns (Experiments 1,2,3). A series of three replicate experiments
ANALYTICAL CHEMISTRY, VOL. 59, NO. 5, MARCH 1, 1987
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/
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Figure 2. 14C03:[14C]-hexadecane ratio vs. time for stored hydrocarbon samples: (0)adsorbed on XAD-2 columns, stored in lab (expt 4); (A)stored as water samples in lab (expt 4).
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Figure 1. Indicator ratios vs. time for stored hydrocarbon samples: sorbed on XAD-2, stored in lab (expts 1,2,3,4)(0); sorbed on XAD2, stored in situ in Ocean (expt 5) (0);stored as water samples in lab (A) (expt 4) (A).(Error bars indicate one standard deviatlon.)
was conducted with standard XAD-2 columns prepared as described above. The standard columns were sealed with plugs made of Teflon immediately after preparation and were stored at room temperature in the laboratory. They were analyzed in duplicate after 0, 1, 3, 10, 32, and 100 days for alkane composition and bacterial numbers. Laboratory Storage of Hydrocarbons and Radiolabeled Hexadecane on XAD-2 Columns (Experiment 4). Radiolabeled hexadecane was added to the input mixture and its degradation into I4CO2was monitored to provide a second, independent measure of bacterial activity. The radiolabeled hexadecane in hexane ([l-'%C]hexadecane,Amersham, 53.6 mCi/mmol hexane) was added to the input mixture to give 0.01 pCi/mL of oil. An initial column was analyzed for alkane composition, ['Vlhexadecane, 'TOz, and bacterial populations. The remaining columns were stored at room temperature, and duplicate columns
were analyzed after 24, 35, and 56 days. To provide a comparison with water samples, 1-L samples were collected in amber bottles from the vat used to load the columns. Three water samples were extracted and analyzed immediately by gas chromatography and concentrationof ['Vlhexadecane and I4CO2. Eight of the water samples were stored sealed at room temperature and were analyzed in duplicate in the same manner as the columns after 3, 6, 11,and 14 days. Nutrients were added to both water and column samples to ensure that nutrient limitation was not a factor in inhibiting bacterial degradation. Nutrient levels were measured in water samples throughout the experiment. Nitrate levels had dropped by the third day, so 5 mL of a 5 g L-' KNOBsolution was added to the remaining water samples. No other adjustments were needed. Nutrient concentrations in columns could not be measured but a fresh solution containing 25 mg L-' nitrate and 2.5 mg L-' phosphate was added to remaining columns at the end of each sample interval. Field Storage of Hydrocarbons on XAD-2 Columns (Experiment 5). A field experiment WBS conducted in which standard columns were hung unsealed at a depth of 1m in seawater from a dock in a small boat marina simulating long-term deployment of in situ sampling equipment. Since the columns were open to the sea, they were exposed to additional bacterial populations, nutrients, and oxygen, so providing a "worst case" experiment for testing the stability of sorbed samples. These columns were analyzed in duplicate for alkane composition and bacterial numbers after 5, 17, and 57 days of deployment. Field Storage of Hydrocarbons on CI8-SilicaGel Columns (Experiment 6). Four standard C18-silica were hung unsealed from a dock in a boat marina at a depth of 1 m; the same conditions existed as described above for XAD-2 columns. They were recovered in duplicate and analyzed for alkane composition and bacterial numbers after 13 and 54 days. To provide a comparison with conventional water samples, 1-Lwater samples from the same input vat used to prepare the columns were stored in the laboratory a t room temperature in amber glass bottles. Duplicate initial water samples and single water samples stored for 3,6,8, 9, and 13 days were analyzed for alkane composition and bacterial numbers.
RESULTS Laboratory Storage of Hydrocarbons on XAD-2 Columns (Experiments 1,2,3). In the three replicate experiments, no evidence of changes in alkane composition or in the ratios indicative of biological degradation were observed over the maximum storage periods of 100 days (Figure 1). Bacterial concentrations in the columns were from lo3 to 104/g throughout the storage period. (No bacteria were found on blank columns.) Laboratory Storage of Hydrocarbons and Radiolabeled Hexadecane on XAD-2 Columns (Experiment 4). The three indicator ratios (pristane: C1,,phytane: C18and U C M C18)clearly indicated bacterial degradation in water samples but not in column
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Figure 4. Indicator ratios vs. time for stored hydrocarbon samples: (0)sorbed on C,,-silica gel, stored in situ in ocean (expt 6): (A)stored as water samples in lab (expt 6).
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Figure 3. Chromatographic traces illustrating comparative degradation of cnde oil in water samples and on XADP columns (expt 4): A, initial sample; 6, column sample after 35 days of storage: C, water sample after 14 days of storage.
samples (Figure 1). Degradation of water samples but not column samples was also demonstrated by changes in [I4C]hexadecane and ‘eo2during storage (Figure 2). The radiolabeled hexadecane in the input mixture samples held in bottles was completely degraded by the eleventh day and 14C02was produced, whereas [14C]hexadecane concentrations on columns remained stable over 56 days of storage and I4CO2was not generated. The contrast in sample stability was also very apparent on the gas chromatograms: stored water samples showed marked degradation after 13 days while column samples were virtually identical with initial samples after long-term storage (Figure 3). Bacteria were present in both column and water samples at similar concentrations (column, 200 to 6.7 X 103/g; water, 60 to 8.3 X 103/mL). Field Storage of Hydrocarbons on XAD-2Columns (Experiment 5). Samples stored on XAD columns open in the ocean showed no signs of biological degradation after 57 days of storage (Figure 1). Bacterial populations observed in columns stored in the ocean were significantly larger than initials after 17 days of storage ((7.2 & 3.3) X 106/g resin ( n = 2) vs. (2.2 f 2.1) X 104/g of resin ( n = 13)), presumably due to additional colonization by bacteria infiltrating from the ocean. Field Storage of Hydrocarbons on C18-Silica Gel (Experiment 6). No evidence of sample degradation, as indicated by changes in alkane composition, was observed when columns packed with CI8-silica gel were stored open in the ocean. Even after 54 days of storage there was no significant change in the various indicator ratios (Figure 4). This preservation effect
occurred despite high concentrations of bacteria on the columns (e.g., 13 days, (2.2 f 1.5 X 106/g resin ( n = 4)). In contrast, marked degradation was observed in water samples taken from the same input vat and stored in the laboratory. Hydrocarbons in the stored water samples were completely degraded after 13 days of storage. The pristane:CI7,phytane:& and UCMC18ratios increased dramatically over the storage period (Figure 4).
DISCUSSION All of our experiments document that hydrocarbon samples are preserved from bacterial attack when on the solid-phase adsorbents XAD-2 and CI6:silicagel. The preservative effect occurs despite the presence of ample oleophilic bacteria populations that, in corresponding water samples, cause complete degradation of hydrocarbons over much shorter storage periods. The mechanism of the preservative effect in solid-phase columns does not appear to be nutrient or oxygen limitation of the bacteria in the columns since ample nutrients and dissolved oxygen were available in two experiments where columns (both XAD-2 and C16:silicagel) were left open in the ocean. We speculate that the preservative effect results from trapping of organics within the adsorbents’ lattice structure. The XAD-2 macroreticular structure has 90-8, holes which are smaller by an order of magnitude than bacteria. The bacteria therefore have access only to hydrocarbons on the outer surface of the resin particles. Similarly, the silica gel used as a substrate for the CIShas a nominal pore size of 90 8,and would similarly sequester hydrocarbons from bacterial attack. In considering these results, it is important to keep in mind the limitations of the study. We worked only with dispersed crude oil, and only at a single, though representative, con-
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centration. However, the preservation effect was replicated many times, under both laboratory and field conditions, and the results were robust. The results are supported by a similar preservation effect noted in the literature for the insecticide fenitrothian sorbed on XAD-2 resin (11, 12).
CONCLUSIONS Both XAD-2 resin and C18-silica gel, when used to concentrate hydrocarbons from water, preserved extracted hydrocarbons from bacterial attack. The preservation of extracted samples was found to be unaffected by differences in dissolved oxygen, nutrients, or size of bacterial population. Safe storage times of columns in the field were a t least 2 months, and in the laboratory at least 3 months. These results contrasted sharply with equivalent stored water samples which showed degradation beginning after a few days and complete degradation after 10-15 days. The preservation effect is encouraging for the future use of solid-phase columns for the extraction of trace organics from water either in situ or in the laboratory. Column samples can be expected to be stable under reasonable conditions of handling and storage for lengths of time up to several months. Samples concentrated on columns can therefore be handled with fewer precautions than ordinary water samples; for instance, they could be mailed to laboratories for analysis without degradation occurring en route. Registry No. XAD-2, 9060-05-3; H20, 7732-18-5.
LITERATURE CITED Green, D. R. Sampling Sea Water for Trace Hydrocarbon Determina tion; National Research Council of Canada publication No. 16472; 1976. Junk, G. A.; Richard, J. J.; Grieser, M. D.; Witiak, D. et ai. J. ChromatOgr. 1974, 99, 745-762. Gallant, R. F., King, J. W.; Levins, P. L,; Piecewicz, J. F. Characterization of Sorbent Resins for Use in Environmental Sampling; - - EPA-600/778-054; 1978. Schatzberg, P.; Adema, C. M.; Thomas, W. M.; Mangum, S. R. Oceans '86 Conference Record; Marine Technology Society: Washington, DC, 1986; pp 1155-1159. Jadamec, J. R.; Kleineberg, G.; Adrick. M. S.; Su, C. H.; Hiltabrand, R. R.; Cutler, J. L., Jr., I n Proceedings of Marine Data Systems International Symposium; New Orleans, LA, April 30-May 2, 1986; Marine Technology Society: 1986; p 127-140. Green, D. R.; Stull, J. K.; Heesen, T. C. Mar. Pollut. Buii. 1986, 17, 324-329. Sander, L. C.; Wise, S.A. Anal. Chem. 1984, 5 6 , 504-510. Jobson, A.; Cook, F. D.; Westlake, D. W. S . Appi. Microbiol. 1972, 2 3 , 1062-1089. National Research Council. I n Oil in the Sea. Inputs, Fates and Effects; National Academy Press: Washington, DC, 1965; pp 32-36. Walker, J. D.; Colwell, R. R. Appi. Environ. Microbioi. 1976, 3 1 , 189- 197. Berkane, K; Caisse, G. E.; Mallet, V. N. J. Chromatogr. 1977, 139, 386-390. Mallet, V. N.; Brun, G. L.; MacDonald, A. N.;Berkane, K. J , Chroma tOQr. 1970, 160, 81-88.
RECEIVED for review July 29,1986. Accepted October 27,1986. This research was supported by the Marine Analytical Chemistry Standards Program of the National Research Council of Canada, and by the Unsolicited Proposal Program of the Canadian Department of Supply and Services.
Determination of Cobalt, Copper, Mercury, and Nickel as Bis(2-hydroxyethy1)dithiocarbamate Complexes by High-Performance Liquid Chromatography Jeffrey N. King and James S . Fritz*
Ames Laboratory-DOE and Department of Chemistry, Iowa State University, Ames, Iowa 50011
Ammonium bls(2-hydroxyethy1)dithiocarbamate (HEDC) is used as a precoiumn derivatiring reagent for the reversedphase HPLC determination of Co( I I), Cu( I I), Hg( I I), and Ni( 11). The metal-HEDC complexes are soluble in water, which eliminates the need to extract them Into an organic solvent prior to analysis. Co( 11), Cu( I I), and Ni( I I)can be determined by direct aqueous InJectiononto a C18 column in the range of 0.005-10.0 mg/L, with a precision of 1.5-3.2%. Detection Is at 405 nm. The Hg( II)-HEDC complex is preconcentrated on an on-line adsorption column prior to HPLC analysis. After preconcentratlon, Hg( I I ) can be determined in the range of 0.02-25 pg/L with a precision of less than 2 % The analysis of spiked electroplating wastewaters showed good agreement with expected values.
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The separation and determination of metal ions by both normal- and reversed-phase HPLC have received a growing amount of attention in recent years. Typically, these methods are based upon the precolumn formation, separation, and subsequent detection of the metal chelates. Several reviews have summarized the work being done in this field (1-3). The disubstituted dithiocarbamates have found extensive use for the separation of metal ions by HPLC. Their ability to form stable chelates with a number of metal ions makes 0003-2700/87/0359-0703$01.50/0
them ideal for this type of application. Separations have been reported by use of both normal-phase (4-10) and reversedphase (11-14) HPLC. Applications have included the analysis of electroplating solutions with a Waters Radial Compression column (15); the automated analysis of wastewater with electrochemical detection (16-19); the determination of trace metals in rice flour and citrus leaves (20);and the determination of low levels of precious metals (21). The dithiocarbamates used thus far for the HPLC separation of metal ions all form water-insoluble metal complexes. When these are used as precolumn derivatization reagents, the metal complexes precipitate as colloidal particles. Thus, the complexes have typically been extracted into an organic solvent such as chloroform prior to the chromatographic separation. The organic extraction can be directly injected onto the separating column when using normal-phase HPLC (9, 10). For reversed-phase HPLC, the organic extract is usually evaporated to dryness and then redissolved in the mobile phase prior to injection (15, 22). Both techniques lengthen the analysis time and increase the possibility of contamination. The direct injection of aqueous sample solutions has been accomplished by incorporating the dithiocarbamate reagent in the mobile phase (16-19, 22). The metal complexes are formed by on-column derivatization prior to the separation. However, detection of the metal complexes is complicated by @ 1987 American Chemical Society
