Anal. Chem. 1084, 56,247-250
In conclusion, the proposed method is very accurate and precise for determination of C1-CB dcohols in gasoline/alcohol fuel blends. The estimated lower detection limits for C1-C3 alcohols and water are 50 and 25 ppm, respectively, injecting 200 pL (max. 100 p L per column) of neat gasohol samples. It requires minimum sample preparation and yields the simultaneous determination of C1-C3 alcohols and water in gasoline/alcohol blends.
ACKNOWLEDGMENT The author thanks R. E. Baker and R. K. Jensen of the Fuels and Lubricants Department for their helpful discussions with this work. Registry No. Ethanol, 64-17-5; methanol, 67-56-1;2-propanol, 67-63-0; tert-butyl alcohol, 75-65-0; water, 7732-18-5.
LITERATURE CITED (1) Pauls, R. E.; McCoy, R. W. J. Chromatogr. Scl. 1981, 79, 558-561. (2) Wong, J. L.; Jaselskis, B. Analyst (London) 1982, 707, 1282-1285.
247
IDurand, J. P.; Petroff, N. Rev. Inst. Fr. Pet. 1982, 37, 575-578.
Chem. Abstr. 1982, 97, 112187a. Sevcik, J. HRC CC, J . H/gh Resolut. Chromatogr. Chromatogr. Commun. 1980, 3 , 166-168. Battiste, D. R.; Fry, S. E.; White, F. T.; Scoggins, M. W.; McWilliams, T. B. Anal. Chem. 1981, 53, 1096-1099. Konopiynski, A.; Siedlecki, A. J. Chem. Anal. (Warsaw) 1980, 25, 777-781. Chem. Abstr. 1980, 95, 45612d. Bjorkqvist, B.; Toivonen, H. J. Chromatogr. 1979, 178, 271-276. Hendrickson, J. G.; Moor, J. C. J. folym. Sci., Part A 1968, 4 , 167-1 88. Smith, W. B.; Kollmansberger, A. J. f h y s . Chem. 1965, 69, 4157-41 61. Hendiickson, J. 0. Anal. Chem. 1968, 40, 49-53. C a m , J.; Gaskill, D. R. Sixth GPC International Seminar Preprlnts, Miami, FL, Oct 1968, 147-157. McComas, D.; Benson, J. Presented at the Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy, Oleveland, OH, 1979; paper no. 3. McKay, V.; Stevenson, R. Presented at the Pittsburgh Conference on Analytical Chemistry and Applied Spectroscopy, Celeveland, OH, 1979; paper no. 98.
RECEIVED for review July 22,1983. Accepted October 3,1983.
Determination of Total Phthalate in Urine by Gas Chromatography Phillip W. Albro,* Satldra Jordan, Jean T. Corbett, and Joanna L. Schroeder National Institute of Environmental Health Sciences, National Institutes of Health, U.S. Public Health Service, P.O. Box 12233, Research Triangle Park, North Carolina 27709
Whlle urlne rarely contalns slgnUlcant quantnles of phthalate diesters, It does contaln a variety of metabolites (phthalate monoesters) Including conjugates. I n the absence of baicterlal action followlng excretlon, the metabolltes retain an Intact phthalate rlng. A procedure for the hydrolysls 6f phthalate esters and metabolltes to free phthallc acld, recovery and esterlflcatlon of the acld, and gas chromatographic quantlflcatlon of the product ester all relatlve to an Internal standard of 4-chlorophthalate has been developed. The measurement limit Is 0.5 nmol of total phthalate/mL of urlne, and the relatlve standard devlatlon Is approximately 1.8% for four or more replicates. The assay Is llnear between 0.5 and 50 nmol/mL urlne, which spans the range of phthalate levels found thus far In human urlne samples. The procedure can also be used to determlne levels of lsophthalate and terephthalate slmultaneously wlth phthalate.
Phthalate esters are used primarily as plasticizers for vinyl and related plastics, with an annual production exceeding a billion pounds (1). Dimethyl phthalate is used topically as an insect repellent (21,bis(Zethylhexy1) phthalate is present in vinyl blood storage bags from which it migrates into stored blood (3-5),and several phthalates are found in medical grade vinyl tubing used for infusions and dialysis therapy (6,7). Two of the major sources of phthalates in the home are vinyl floor tiles and electrical insulation, from which the plasticizer slowly volatizes as the products age (2). Although most phthalates have a very low order of acute toxicity, reports that high-level chronic exposure to bis(2ethylhexyl) phthalate causes liver cancer in rats and mice (8) while several phthalates including bis(2-ethylhexyl) and diThis artlcle not subject to
n-butyl (the two most widely used phthalate plasticizers) cause testicular atrophy in rats (9)have stimulated a great deal of concern about the potential hazard of these compounds to humans. Phthalate diesters have been found in a variety of animal tissues (10,II)and food (12,13)but are very rapidly cleared from human blood after exposure (14). In general, intact diesters of the more common plasticizers are not found in urine (except dimethyl phthalate (15)). In most cases only hydrolyzed and oxidized metabolites occur in urine of mammalian species (16);however, mammals appear to be unable to metabolize the phthalate ring system, which is intact in all of the identified metabolites ( 17 ) . Metabolites of phthalate diesters are eliminated from the body in both feces and urine (18),as a mixture of free and glucuronide-conjugated acidic forms in the case of all animals studied extept rats (If?),which do not form conjugates of bis(ethylhexy1) phthalate metabolites. Therefore analysis of clinical samples for phthalate diesters will not effectively reflect exposure. Since a large number of different metabolites are excreted (17),it would be most useful to have a single meaburemknt of total phthalate as an indirect indicator of level of exposure. The ratio of urinary to fecal excretion of phthalate metabolites appears to be reasonably constant for a given plasticizer in a given species but may differ for different plasticizers (15). Therefore, determination of total urinary phthalate can only serve as a relative indicator of exposure and not as an absolute measure. The classical procedure for determination of phthalate in urine (19) involved hydrolysis to phthalic acid, oxidation of interferences with nitric acid, and precipitation of phthalic acid as the lead salt for gravimetric determination. This procedure has obvious limitations in terms of specificity and sensitivity. The method described in the present paper was
U S . Copyright. Publlshed 1984 by the American Chemical Society
248
ANALYTICAL CHEMISTRY, VOL. 56, NO. 2, FEBRUARY 1984
developed to facilitate the measurement of phthalate at levels between 0.5 and 50 nmol/mL of urine. EXPERIMENTAL SECTION Urine is collected and stored frozen over a crystal of thymol in a glass container equipped with a Teflon-lined screw cap. All such glassware must be thoroughly rinsed with phthalate-free acetone before use. Specific gravities were measured with an American Optical temperature-compensated TS Meter Series B (American Optical Co., Keene, NJ). Distilled, deionized water was passed through activated charcoal to ensure freedom from phthalate contamination and to provide a procedural blank reference. Gas chromatography was performed with a 3 m x 2 mm i.d. stainless steel column packed with 10% OV-25 on 100/120 mesh Gas Chrom Z, isothermal at 180 "C with a helium flow rate of 37 mL/min. Injection port and hydrogen flame ionization detector temperatures were 250 and 270 "C, respectively. confirmation may be accompLshed with a 2 m X 2 mm column packed with 10% OV-225 on 100/120 mesh HP Chromosorb W (Applied Science Laboratories, Deerfield, IL) at 175 OC, 30 mL of helium per min. Interference is likely on less polar liquid phases. Chemicals used in this work include the following: phthalic acid, dimethyl isophthalate, and terephthalic acid from Analabs, North Haven, CT; suberic acid, diethyl, dibutyl, and bis(2ethylhexyl) phthalate, and N-methyl-N'-nitro-N-nitrosoguanidine from Aldrich Chemical Co., Milwaukee, WI; 14% BF3-methanol reagent, Applied Science; thymol, di-n-octyl phthalate, dimethyl phthalate, diethyl ether, n-hexane, and sodium hydroxide (5 N solution),Fisher ScientificCo., Raleigh, NC; and 4-chlorophthalic acid from Sigma Chemical Co., St. Louis, MO. All other organic solvents were HPLC grade glass-distilled. Bis(2-ethylhexyl) phthalate labeled with I4C in the carbonyl group was prepared as described previously (20). Acidic alumina 90 (Merck) was activated at 130 "C in an oven overnight before use. Total urinary metabolites of bis(2-ethylhexyl) [7-14C]phthalatewere produced in CD-1 mice as described previously (16). Internal Staadard. An internal standard for quantitation must be added to the urine prior to hydrolysis of extraction. Either suberic acid (1,6-hexanedicarboxylic acid), which has been recommended previously (17), or 4-chlorophthalic acid may be used as internal standard, at a level of 20 nmol to 1 pmol of standard/mL of urine. The standard chosen is first dissolved in methanol to give a 0.02 M solution. The standard solution is then added to the urine with a Kirk micropipet and mixed well. Base Hydrolysis. To 10 mL of urine of known specific gravity in an 18-mL screw cap test tube are added 10 pL of internal standard and 2.5 mL of 5 N NaOH. The tube is capped (Teflon-lined screw cap) and the mixture is mixed and heated at 95 & 5 OC for 90 min. After cooling, the solution is transferred to a 60-mL separatory funnel equipped with a Teflon stopcock. Slowly and cautiously, 2.5 mL of concentrated (approximately 12.5 N) HC1 is added, mixed, and allowed to cool. The suspension is extracted at least twice and preferably four times with 20-mL portions of diethyl ether. Combined extracts are dried over anhydrous sodium sulfate, filtered, and evaporated to dryness at 40 "C with a rotary evaporator. Small portions of dichloromethane may be added and evaporated to remove residual moisture. If the urine is for some reason anticipated to contain significant amounts of bis(2-ethylhexyl) phthalate, the above hydrolysis procedure will not be adequate. Instead it will be necessary to employ 2 N NaOH or KOH in 50% aqueous ethanol as described previously (20). Esterification. The residue from the evaporation flask after hydrolysis and ether extraction is dissolved in 2 mL of 14% BF3-CH,0H reagent and transferred to another 18-mLscrew cap test tube (Teflon-lined screw cap). The tube is capped and heated at 95 f 5 "C for 1 h. After the tube cools, 1mL of n-hexane and 12 mL of water are added, mixed vigorously, and centrifuged to obtain a complete phase separation. Most of the hexane (upper) layer is transferred into a test tube. The lower phase is reextracted with 3 mL of diethyl ether and centrifuged and the ether extract combined with the previous hexane extract. The combined extract is evaporated nearly to dryness under nitrogen and then diluted to 1 mL with n-hexane. If there are only a small number of samples to be processed,
it may be more convenient to perform the esterification with ethereal diazomethane (21). However, a definite excess of diazomethane must be assured, and the coloration of the extract will mask the yellow of the diazomethane. A variety of esterification procedures were tested as to their ability to convert phthalic acid to dimethyl phthalate, with the constraint that applicability to potentially large numbers of samples precluded reflux conditions. Acid-catalyzed esterification with methanolic HC1 or H2S04(22,23), esterification catalyzed by thionyl chloride (241, the use of dimethylformamide dimethyl acetal (W), acylation of tetramethylammonium salts or silver salts with methyl iodide (26,27),and transesterification of trimethylsilyl esters (28) were attempted on mixtures of suberic and phthalic acids and the results compared to those obtained with diazomethane and BF3-CH30H. In some cases 2,2-dimethoxypropane stabilized with MezSO was used as a water scavenger in the acid-catalyzed reactions (29). Cleanup. Pasteur pipets plugged with glass wool are packed dry with 0.5 g of acidic alumina that has been activated at 130 "c overnight. A 0.5-cm layer of granular, anhydrous sodium sulfate is placed above the alumina, and the packing held in place by a second glass wool plug. The hexane solution of esterification products is loaded onto the column and washed with 5 mL of n-hexane:dichloromethane,3:2 (v/v). This wash may be discarded. The desired products (dimethyl esters) are eluted with dichloromethane:acetone,9 1 (v/v), collecting the first 2 mL to elute. Aliquots (3 pL) of this fraction are taken for gas chromatographic analysis as described above. Recovery may be checked by adding 1 clmol of diethyl phthalate to this solution as a second internal standard. Alternative Procedure for Isophthalate and Terephthalate. In some cases it may be desired to measure not only o-phthalate but also isophthalate (m-) and terephthalate 9-). These isomers, especially terephthalate, are, as the free acids, so nearly insoluble in diethyl ether that it will be impossible to extract them in reasonable recovery from the hydrolysis products. They may be recovered after acidication with HCl (the pH must be one or below) by chromatographic stripping. Columns (e.g., burets plugged with glass wool and equipped with Teflon stopcocks) of 1 cm i.d. are packed to a depth of 9.5 cm (ie., 7.5 mL bed volume) with 20-50 mesh XAD-2 resin as a slurry in methanol. Before use, the resin must be Soxlet-extracted with methanol and never subsequently be allowed to dry out. The packed columns are topped by a 2-cm layer of Sea Sand (to prevent flotation of the resin) and then washed by elution with 75 mL of distilled, charcoal-treated water followed by 10 mL of 0.1 N aqueous HCl. The hydrolyzed and acidified urine is passed through the column followed by 50 mL of 0.1 N aqueous HC1. When the diluted HC1 reaches the top of the sand, 2 mL of ethyl acetate is added to form an interface, followed by methanol. Collection begins as the ethyl acetate begins to elute, and a total fraction of 25 mL (2 mL of ethyl acetate plus 23 mL of methanol) is collected. Solvent may be removed by rotary evaporation at 40 "C. RESULTS AND DISCUSSION Recovery. Twenty samples of urine containing between 0.6 and 25 nmol of phthalate per mL gave a mean recovery of 77.2 f 16.1% (standard deviation) relative to diethyl phthalate added just prior to GC if the acidified hydrolysis mixture was extracted only twice with diethy ether. This could be tolerated when 4-chlorophthalate was used as internal standard, as there was no statistically significant difference in paired comparisons between its recovery and the recovery of phthalic acid. However, suberic acid had a higher partition coefficient (3.6 vs. 1.0) than phthalic acid under these conditions, and it was necessary to extract with ether four times (giving greater than 93% recovery of both acids) when suberic acid was used as internal standard. The variability of recovery was much less even with only two ether extractions (RSD = 7.7% of the mean) when six aliquots of the same urine sample, containing 11.9 A 0.87 nmol of phthalate/mL, were processed. Thus the particular composition of a given urine sample can influence the recovery of phthalic acid, and for this reason the more structurally
ANALYTICAL CHEMISTRY, VOL. 56, NO. 2, FEBRUARY 1984
Table I. Yield of Dimethyl Phthalate from Phthalic Acid in BF,-CH,OH Reagenta
Table 11. Gas Chromatographic Parameters for Analytes and Reference Compounds
min at 95 "C % yieldb min at 95 "C % yieldb 15 20
47.1 82.2
30 60
99.5 99.9
a One-micromole samples of phthalic acid heated in capped tubes with 2.0 mL of 14% BF,-CH,OH ( 3 1 ) for the times indicated. After the reaction was stopped with 12 mL of water, 1 pmol of dimethyl suberate was added, the products were extracted with hexane and ether, and analysis was performed by gas chromatography as discussed in the text. Percentage of theoretical yield of 1 pmol of dimethyl phthalate. Monomethyl phthalate not included.
similar 4-chlorophthalate may be the more reliable internal standard. Moreover, urine samples may occassionally contain small amounts of suberic acid as a metabolite of suberate plasticizers or longer-chain dicarboxylic acids (30). Hydrolysis. Total urinary metabolites of bis(Zethylhexy1) [7-14C]phthalate, which included both free and glucuronidewere converted quantitatively to [7conjugated forms (I6), 14C]phthalic acid under the described conditions, as determined by both thin-layer chromatography and radio-HPLC (I7,20). Unhydrolyzed phthalate esters were not detectable when urine was fortified with dimethyl, diethyl, di-n-butyl, or di-n-octyl phthalate and subjected to the described conditions. Only bis(Zethylhexy1) phthalate showed enough steric hindrance to be a problem, being only 60% hydrolyzed in 90 min in 1N aqueous NaOH. However, as discussed above this hydrophobic diester is not normally found in human urine unless inadvertently introduced as a contaminant after collection of the urine. Esterification. Acylation of phthalic acid salts was unsatisfactory as an esterification procedure because of the extreme insolubility of either TMAH or Ag salts in the presence of either dimethylformamide or dimethydacetamide (26). Transesterification of trimethylsilyl esters was ineffective-no peaks were seen during gas chromatography. The dimethylformamide dimethyl acetal procedure (25) was attempted in either acetonitrile or chloroform, but was unsatisfactory. Dimethyl esters did form, but decomposed to unidentified products, and a quantitative yield could not be achieved in spite of the use of a variety of times and temperatures. Artifacts (extra peaks) were formed with thionyl chloride, and the use of dimethoxypropane resulted in the conversion of phthalic acid to phthalic anhydride which was recovered (nonquantitatively) as such. Simple acid-catalyzed esterification in the absence of a water scavenger was effective, but appeared to reach an equilibrium point at 88-93% esterification. Only diazomethane and the BF3-CH30Hreagent appeared capable of quantitative esterification of o-phthalic acid under the conditions tested. Dimethyl phthalate has sufficient solubility in aqueous methanol that hexane alone would not give complete extraction and diethyl ether was required. However, the preextraction with hexane greatly reduces the amount of water taken up by the ether, eliminating the need for an additional drying step. Table I shows the yieId of dimethyl phthalate from phthalic acid in the reaction with BF3-CH30H as a function of time. The reaction is complete after 30 min a t 95 OC, and the products are stable. Preformed dimethyl phthalate was treated with BF3-CH30H for 60 min a t 95 "C with no detectable loss; however, it is essential that all residual water be evaporated following ether extraction of the hydrolysis produds and before addition of the BF3-CH30H. The cleanup on alumina removes mainly highly polar material that would otherwise build up on the gas chromatographic column.
249
compound dimethyl suberate dimethyl phthalate dimethyl isophthalate dimethyl terephthalate dimethyl 4-chlorophthalate diethyl phthalate
OV-25, OV-225, 180 "C 175 "C R M R ~R R T ~R I ~ RRT RI 0.993 1,000 1.049 0.971 0.959
0.553 1.000 1.055 0.926 1.535
1728 1879 1892 1859 1987
0.519 1.000 0.953 0.859 1.524
1919 2095 2082 2054 2209
1.258 1.593 1996 1.490 2204
Relative molar response of the hydrogen flame ionization detector to the compound specified, compared to its response to the corresponding molar quantity of dimethyl Corrected retention phthalate, in terms of peak areas. time of the compound specified, divided by corrected Kovats retention retention time of dimethyl phthalate. index ( 3 2 ) .determined relative to a series of a-alkanes.
Figure 1. Sample gas chromatograms: (A) standards on OV-25; (B) urine fortified with phthalic and chlorophthalic acids, complete procedlrre, OV-25 % (C) standards on OV-225; (D) human urine with suberic acid internal standard, diethyl phthalate added as secondary standard just before injection (phthalate level found, 31 nmol/mL OV-25); (E) same sample as D, on 011-225; (F) charcoal treated water, as for D; peak indentifications, (a)dimethyl suberate, (b) dimethyl phthalate, (c) dimethyl 4-chlorophthalate, (d) diethyl phthalate. For operating conditions see text. G a s Chromatography. Table I1 lists relative retention times (corrected for dead space), Kovats retention indexes (32), and relative molar detector responses (dimethyl phthalate response set equal to 1.000) under the recommended chromatographic conditions. The peaks are sufficiently symmetrical (Figure 1)that either height times width at half height or an electronic integrator can be used to determine peak areas. Retention times remained constant to within 1.5% over a 5-week period of constant use of the OV-25 column, suggesting that peak heights could be used for quantitation. However, base line resolution is not always achieved, and peak areas @reconsidered more reliable. Quantitation. No peaks were seen a t the dimethyl phthalate position when 10-mL samples of charcoal-treated water were fortified with internal standard and run through the entire procedure. Although bis(2-ethylhexyl) phthalate is a common laboratory contaminant and found in most organic solvents (33),it is resistant to transesterification and will not cause interference in this procedure as long as it does not contaminate the NaOH or test tubes used in the hydrolysis. Di-n-butyl phthalate, also commonly found as a trace contaminant of organic solvents, may transesterify in BF,-
250
9
ANALYTICAL CHEMISTRY, VOL. 56, NO. 2, FEBRUARY 1984
CH,OH, so must be absent from the esterification reagent and from the diethyl ether used to extract the hydrolyzed urine. Its introduction after the esterification step will not cause interference. In order to test for the possibility of dechlorination of the 4-chlorophthalate by NaOH, water samples were spiked with highly purified dimethyl 4-chlorophthalate (purified by preparative HPLC on Spherisorb S5CN by elution with nhexane:methyl tert-butyl ether, 85:15) and put through the described complete procedure. No dimethyl phthalate was detected (less than 0.001%). Since no samples of human urine have been found, in our laboratory, to be aompbtely free of phthalate, urine samples were divided into geveral aliquots for analysis before and after fortification wjth 20 nmol sf ethylhexyl phthalate (a major urinary metabolite of bis(ethylhexy1) phthalate in humans (14)), per mL of urine. After subtracting the apparent amounts of phthalate in the unfortified samples from that found after fortifjcation, the residual was 19.5 f 1.4 (standard deviation) nmol/mL ( N = 6) with suberate as internal standard. This improved considerably to 20.1 f 0.35 (N = 4) with 4-chlorophthalic acid as internal standard, probably because dimethyl suberate was not base line resolved from other urine constituents under these chromatographic conditions. Thus the advantage of a relative molar response equivalent to that of dimethyl phthalate is more than offset by the reduced precision associated with the use of suberate as internal standard. ' We have never observed interference with the 4-chlorophthalate peak from any sample of human urine, although no guarantee that such will never occur can be given. Mixtures of dimethyl phthalate and dimethyl 4-chlorophthalate at molar ratios from 0.025 to 1.25 were made up in methylene chloride such that the concentration of 4-chlorophthalate was 0.5 pmol/mL. These mixtures were analyzed on OV-25, and the ratios of peak areas (phthalate/chlorophthalate) plotted against molar ratios. A regression line through the data points had the equation: [(pmol of phthalate)/(pmol of chlorophthalate)] = [0.9597 X (phthalate peak area/chlorophthalate peak area)] 0.0006. The linear correlation coefficient was 0.9995 (five replicates). Thus one should use 200 nmol of 4-chlorophthalate as internal standard in order to measure down to 0.5 nmol of total phthalate/mL of urine in a 10-mL sample. The slope of 0.9597 corresponds to the relative molar response of the flame detector to dimethyl chlorophthalate. If the urine was obtained from persons known to be occupationally or otherwise exposed to phthalates, it may be advisable to use 1 pmol of internal standard. Precision. To check the replicability of the procedures, urine specimens previously found to contain apparent levels of total phthalate at the low, middle, and high regions of the applicable range of the assay were divided into sets of replicates for analysis. Levels of total phthalate observed were 0.79 f 0.073 nmol/mL (N = 5, RSD = 9.3%), 12.2 f 0.22 nmol/mL (N = 5, RSD = 1.8%),and 53.1 f 0.99 (N = 6, RSD = 1.9%). The number of replicates was dictated by the amount of urine available; a larger number of replicates might have lowered the RSD for the low level specimen. Samples fortified with a mixture of phthalic and terephthalic acids and processed by the alternate procedure (XAD-2 rather than diethyl ether to recover the products after treatment with NaOH) gave a mean recovery of 96.9 f 5.0% of terephthalate relative to phthalate. Since isophthalate is
-
intermediate between the two in aqueous solubility, the alternate procedure should be applicable to all three. These three isomers have been separated (as pure standards of the dimethyl esters) on an OV-101 glass capillary column by Friocourt and co-workers (34),but this liquid phase (and other nonpolar dimethyl polysiloxane phases) shows interference with the dimethyl phthalate peak frpm normal urine constituents. Moreover, it fails to resolve dimethyl phthalate from dimethyl suberate in our hands. Over 200 distinct samples of human urine have been processed by the described technique; in no case did the material cochromatographing with dimethyl phthalate on OV-25 fail to cochromatograph with dimethyl phthalate on OV-225, which differs from OV-25 in selectivity (a.g., dimethyl 4chlorophthalate and diethyl phthalate reverse elution order on the two phases). Thus we consider phthalates to be ubiquitous in buman urine, most probably'as a result of environmental exposure. This conclusion must remain speculative, however, until nonchromatographicconfirmation of the phthalate identification is available for a large number of samples.
LITERATURE CITED (1) Pierce, R. C.; Mathur, S. P.; Williams, D. T.; Boddington, M. J. Phthalate Esters in the Aquatic Environment"; NRCC No. 17583; Envlronmental Secretariat, NRCCKNRC: Ottawa, 1980. (2) Fishbein, L; Albro, P. W. J . Chromatogr. 1972, 7 0 , 365. (3) Marcel, Y. L.; Noel, S. P. Lancet 1970, 1 , 35. (4) Jaeger, R. J.; Rubin, R. J. New Engl. J . Med. 1972, 278, 1114. (5) Kim, S. W.; Petersen, R. V.: Lee, E. S. J . Pharm. Scl. 1976, 670. (6) Glbson, T. P.; Briggs, W. A.; Boone, E. J. J . Lab. Clin. Med. 1976, 6 7 , 519. ( 7 ) Ono, K.; Tatsukawa, R.; Waklmoto, T. JAMA, J . Am. Med. Assoc. 1975, 234, 948. (8) Kluwe, W. M.; McConnell, E. E.: Huff, J. E.; Haseman, J. K.; Douglas, J. F.; Hqrtweli, W. V. EHP, Environ. Health Perspect. 1982, 4 5 , 129. (9) Gray, T. J. 6.; Rowland, I.R.: Foster, P. M. D.; Gangolli, S. D. Toxlcol. Len. 1982, 1 1 , 141. IO) Nazir, D. J.; Alearaz, A. P.; Bierl, 6.A,; Beroza, M.; Nair. P. P. Biochemistry 1971, 10, 4228. 11) Mes, J.; Coffin, D. E.; Campbell, D. S. Bull. Environ. Contam. Toxlcol. 1974, 12, 721. 12) Tomlta, I.; Nakamura, Y.; Yagi, Y . Ecotoxicol. Environ. Saf. 1977, 1 , 275. 13) Burns, B. G.; Musiai, C. J.; Uthe, J. F. J . Assoc. OM. Anal. Chem. 1981, 6 4 , 282. (14) Peck, C. C.;Albro, P. W. EHP, Environ. Health Perspect. 1962, 45, 11
(15) Albro, P. W.; Moore, B. J . Chromatogr. 1974, 9 4 , 209. (16) Albra, P. W.; Corbett, J. T.; Schroeder, J. L.; Jordan, S.; Matthews, H. B. EHP, Environ. Health Perspect. 1962, 45, 19. (17) Albro, P. W.; Jordan, S. T.; Schroeder, J. L.; Corbett, J. T. J . Chromatogr. 1982, 2 , 130. (18) Klune, W. M. EHP, Environ. Health Perspect. 1982, 4 5 , 3. (19) Shaffer, C. B.; Carpenter, C. P.; Smyth, H. F., Jr. J . Ind. Hyg. 1945, 2,130. (20) Albro, P. W.; Thomas, R.; Fishbein, L. J . Chromatogr. 1973, 7 6 , 321. (21) Fales, H. M.; Jaouni, T. M.; Babashak, J. F. Anal. Chem. 1973, 4 5 , 2302. (22) Fischer, E. Ber. Dtsch. Chem. Ges. 1906, 3 9 , 2896. (23) Rogozinski, M. J . Gas Chromatogr. 1964, 2 , 328. (24) Gee, M. Anal. Chem. 1965, 3 7 , 926. (25) Vorbruggen, H. Angew. Chem., Int. Ed. Engl. 1963, 2 , 211. (26) Greeley, R. H. J . Chromatogr. 1974, 8 8 , 229. (27) Gehrke, C. W.; Goeriitz, D. F. Anal. Chem. 1963, 3 5 , 76. (28) Mamer, 0.A.; Gibbs, B. F. Clln. Chem. (Winston-Salem, N . C . ) 1973, 19, 1006. (29) Simmonds, P. G.; Zlatkis, A. Anal. Chem. 1965, 3 7 , 302. (30) Mortensen, P. B.; Gregersen, N. Blochlm. Biophys. Acta 1982, 710, 477. (31) Morrison, W. R.; Smith, L. M. J . LipldRes. 1964, 5 , 600. (32) Kovats, E. Helv. Chim. Acta 1958, 4 1 , 1915. (33) Ishida, M.; Suyama, K; Adachl, S. J . Chromatogr. 1960, 189, 421 (34) Friocourt, M. P.; Berthou, F.; Picart, D.; Dreano, Y.; Floch, H. H. J . Chromatogr. 1979, 172, 261.
RECEIVED for review July 8, 1983. Accepted October 26, 1983.
