Biological Monitoring for Pesticide Exposure - American Chemical

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16

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New Approaches to Analysis of Organophosphate Metabolites in the Urine of Field Workers C. P. Weisskopf and J. N. Seiber Department of Environmental Toxicology, University of California, Davis, CA 95616 A procedure is described for the analysis of urinary dialkyl phosphate and thiophosphate metabolites using disposable extraction cartridges, large bore capillary gas chromatography (GC) columns, and the methylating reagent trimethylanilinium hydroxide. Although field worker monitoring for organophosphate pesticide exposure by measurement of urinary metabolites can have several advantages, alternate methods are frequently favored, in part due to difficulties in the analysis of these compounds. The described method overcomes many of the obstacles of conventional approaches to urinary metabolite analysis. Metabolite extraction is rapid, requiring at most 15 minutes per sample. Derivatiza tion takes place in the GC injector block, and capillary columns lead to improved separations for all compounds. Five urinary metabolites are detectable at concentrations of 2 to 10 ng/ml (ppb) urine. There has been a long-standing interest in the estimation of human exposure to organophosphates, particularly since these compounds are widely used and are the most frequent cause of pesticide related occupational illnesses. Upon entering the body, most organophosphorus pesticides may be metabolized to yield one or more of the six common dialkyl phosphates shown in Table I. Quantities of metabolites excreted in the urine have been found to correspond to the pesticide dose (1-3). Measurement of these urinary metabolites can thus provide a basis for comparison of exposure in a study population. Measurement of intact pesticides or metabolites in biological samples may give a better estimation of the amount of material actually entering the body than external measurement methods. Analysis for principle organophosphate pesticide alkyl or aryl leaving groups can yield information on the identity of the pesticide to which the subject was exposed, since few pesticides share the same leaving group. However, this requires the development of an analytical method specific for each compound, and is thus not amenable to a multi-residue approach. The dialkyl phosphates in Table I represent possible metabolites of approximately 75% of the organophosphate pesticides in the EPA's Pesticides and Industrial Chemicals Repository. 0097-6156/89/0382-0206$06.00/0 ° 1989 American Chemical Society

Wang et al.; Biological Monitoring for Pesticide Exposure ACS Symposium Series; American Chemical Society: Washington, DC, 1988.

16.

WEISSKOPF AND SEIBER

Organophosphate Metabolites in Urine

207

Table I. Common dialkyl phosphates

DMP DEP DMTP DETP DMDTP DEDTP

0,0-Dimethyl phosphate 0,0-Diethyl phosphate 0,0-Dimethyl phosphorothionate 0,0-Diethyl phosphorothionate 0,0-Dimethyl phosphorodithioate 0,0-Diethyl phosphorodithioate

The primary disadvantage to the measurement of urinary dialkyl phosphates lies i n the difficulty of the analysis. These compounds are highly water soluble, ionized i n urine, and must be derivatized to be sufficiently volatile for gas chromatographic (GC) analysis. Although there are many published procedures for extraction and analysis of urinary dialkyl phosphates (3-8). the methods remain tedious and cumbersome. The majority rely on liquid/liquid extraction of the metabolites, derivatization with diazoalkane reagents, and analysis by packed column GC. Metabolite extraction is often incomplete or inconsistent (4), and it has been difficult to get adequate chromatographic resolution of all metabolites both from each other and from other constituents of urine (5-7). Diazoalkanes have frequently been used for alkylation of dialkyl phosphates and thiophosphates prior to G C analysis. Diazoalkane reagents such as diazoethane and diazopentane give an isomer mixture of thionate and thiolate esters, formed i n irregular proportions, when the metabolites D M T P or D E T P are derivatized (8). Analysis of both isomers may be required, resulting i n increased analysis time and poorer detection limits. When coupled with the hazards of the preparation and use of diazoalkanes, this makes the use of an alternative derivatizing reagent desirable (4). Trimethylanilinium hydroxide ( T M A H ) is a methylating reagent frequently used in clinical tests for barbiturates. A t elevated temperatures, provided i n this case by the G C injector block, T M A H is able to methylate dialkyl phosphates. Dale et al. (9) first reported the use of T M A H for the hydrolysis of intact organophosphates and methylation of hydrolysis products. Miles and Dale (10) followed with a study that included the use of T M A H for the injector block methylation of DMDTP. Use of T M A H for the methylation of D M T P in the extracts of urine of exposed workers was reported by Moody et al. (U_). The present study was designed to overcome some of the complexities of previous methods by developing a simpler and faster analytical procedure through the use of disposable extraction cartridges, injector block derivatization, and quantification using large bore capillary G C columns. Experimental Section Solvents and Reagents. Trimethylanilinium hydroxide ( T M A H ) was obtained from Pierce Chemical Co, Rockford, IL, under the trade name MethElute, a 0.2 M solution of T M A H i n methanol. Handeling precautions for this mixture are the same as those for methanol. Solvents were pesticide residue grade from J. T. Baker, Phillipsburg, NJ. Salts and acids were analytical reagent grade from Mallinckrodt, Inc., Paris, K Y . Extraction Apparatus. Disposable cyclohexyl extraction cartridges with a 6 ml reservoir containing 1 g sorbent were used (Analytichem International, Harbor City, CA). A ten-place vacuum manifold was used for cartridge elution (J. T. Baker Co, Phillipsburg, NJ). Sufficient vacuum (approximately 100 mm Hg) was


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BIOLOGICAL MONITORING FOR PESTICIDE EXPOSURE

applied to provide for a flow rate of 5 ml/min of eluting solution through the cartridges. Preparation of Standard Solutions. Analytical grade standards of the six metabolites in Table I were obtained from the EPA's Pesticides and Industrial Chemicals Repository. Stock 1 mg/ml solutions of the individual dialkyl phosphates in methanol were prepared and stored at 4°C in the dark. Analytical mixed standards of all six metabolites in acetone (0.015 to 1.5 ng//il) were prepared daily in volumetric flasks, and brought to final volume after the addition of T M A H immediately prior to G C injection. T M A H was added to the standard solutions in the proportion of 50 /il TMAH/ml solution. Chromatographic Conditions. Apparatus: The instrument used was a Tracor model MT 220 equipped with a flame photometric detector (FPD) operated in the phosphorus mode. The column was a 30 m x 0.5 mm ID large bore DB-1701 capillary column, with an 86% dimethyl- and 14% cyanopropylphenyl- polysiloxane bonded stationary phase and a 1.0 /xm film thickness (J & W Scientific, Folsom, CA). Gas flows were: helium carrier gas 7 ml/min; helium detector make-up gas 40 ml/min; air 90 ml/min; and hydrogen 60 ml/min. Operating temperatures were: injector, 300°C.; column, 110°C.; and FPD and FPD transfer line, 170°C. Quantification of the methylated metabolites was performed by external standard calculations based on peak areas using a Hewlett Packard 3390 A electronic integrator. 1-3 /il aliquots of the mixed standard solutions or urine extracts were injected. Standard curves were prepared from duplicate injections of mixed standards at a minimum of three concentrations. Determination of Dialkyl Phosphates in Urine. The preparation of samples for analysis is outlined in Figure 1. An aliquot (4 ml) of urine was pipetted into a 15 ml test tube. Granular ammonium sulfate (ca 1.8 g) and 4 drops 70% acetic acid were added, and the mixture vortexed for 30 sec. The cyclohexyl extraction cartridges were prewashed with one column volume (ca 6 ml) each of acetone, water, and ammonium sulfate-saturated water. The urine sample was then passed through the cartridge. The cartridges were washed with 2 ml of a solution of 20% acetone in hexane, and the wash discarded. The vacuum was increased to about 600 mm Hg, and the cartridges were aspirated for 5 minutes. The cartridges were then placed into 15 ml graduated centrifuge tubes and eluted with acetone to a volume of 1.4 ml by gravity flow or the application of positive pressure to the extraction cartridge using a pipette bulb. Immediately prior to GC injection ca 0.4 g powdered anhydrous sodium sulfate and 100 /xl T M A H was added to the acetone eluate followed by brief vortexing. Results and Discussion The procedure reported here was capable of analyzing DEP, DMTP, DETP, DMDTP, and DEDTP. It is unsuitable for DMP due to interferences caused by the presence of inorganic phosphate. The effect of various salts on the recovery of dialkyl phosphate metabolites was tested in developing this procedure. Salts were added to urine at amounts slightly in excess of that needed to produce a saturated solution, allowing for reproducible results without expending much time in weighing. Salt addition usually improved recovery of metabolites from fortified urine samples, with sulfate salts giving better results than either acetate or chloride salts (Table II). A saturated solution of ammonium sulfate was found to be the best of those tested.


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4 ml Urine add:

CH Cartridge ca 1.8 g (NH ) S0 4 drops 70% HOAc 4

2

wash: acetone water salt saturated water

4

Vortex 30 seconds Add Urine to Washed Cartridge wash: 2 ml 20% acetone in hexane aspirate: 5 minutes Column + Retained Metabolites acetone elution to 1.4 ml Acetone Eluate + Metabolites add:

ca 0.4 g Na^O, 100 Ml TMAH

Gas Chromatography Figure 1. Flow diagram for the analysis of dialkyl phosphates in urine.


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Table II. Effect of salt addition on metabolite recovery, normalized for ammonium sulfate

grams added

salt

none sodium sulfate sodium chloride ammonium sulfate ammonium acetate

-

1.1 1.4 1.7 4.0

DEP

0.1 0.9 0.4 1.0 0.0

normalized recovery DMTP DMDTP DETP DEDTP

0.1 1.0 0.7 1.0 0.0

0.1 0.8 0.7 1.0 0.2

0.2 1.0 0.8 1.0 0.4

0.5 0.9 1.0 1.0 0.5

The p H of the urine was also found to have a large effect on recoveries (Table III). Recoveries dropped when the urine was made more basic than the p H of 5 of the unmodified urine. Recoveries improved when acetic acid was used to adjust the p H to approximately 4, and declined when hydrochloric acid was used to reduce the p H further. Acetic acid was used routinely as the p H modifier. Table III. Effect of p H modification on metabolite recovery, normalized for acetic acid

modifier

sodium hydroxide none acetic acid hydrochloric acid hydrochloric acid

pH

DEP

7 5 4 3.5 2.5

0.5 0.9 1.0 0.4 0.2

normalized recovery DMTP DMDTP DETP DEDTP

0.5 0.8 1.0 0.8 0.6

0.6 0.9 1.0 0.7 0.5

0.6 0.9 1.0 0.8 0.4

0.6 0.9 1.0 0.8 0.4

A hexane/acetone wash followed by aspiration was found to eliminate most of the water held by the cartridges, as well as removing some interfering compounds that reduced the efficiency of metabolite derivatization. Subsequent elution with 1.4 ml acetone removed more than 9 5 % of the metabolites retained on the cartridges. Samples could then be analyzed directly, eliminating the possibility of evaporative loss of the compounds or expenditure of time in reducing solvent volumes, both drawbacks of prior methods which used solvent extraction rather than extraction with solid phase cartridges. The efficiency of T M A H as a methylation reagent was tested using a variety of solvents, sample extracts, and injector temperatures. Derivatization yields were estimated by comparing molar phosphorus response of the methylated metabolites on the flame photometric detector with the response of a trimethyl phosphate standard. Derivatization yields were highest when the solvent was acetone and the injector block of the G C was above 250°C. The methylation yield of D M P could not be determined due to interference from inorganic phosphate. Both compounds are methylated to produce trimethyl phosphate (Figure 2), making T M A H unsuitable for analysis of D M P without prior removal of inorganic phosphate. Approximately 6 0 % of the D E P and 9 5 % of the four other metabolites in acetone are methylated. When the solvent was an acetone extract


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I

0

1 2

1 4

1 6


1 1 1 r 8 10 12 14

i

0

1 2

1 4

1 6

1 1 1 r8 10 12 14

TIME (minutes)

Figure 2. Gas chromatogram of dialkyl phosphates using FPD detection, a) reagent blank; b) dialkyl phosphate standards, 0.015 ng each except DEP, 0.03 ng; c) human urine with no detectable metabolites; d) human urine fortified at 10 ppb each metabolite.


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of urine, DEP derivatization was reduced to 40%; the other compounds were unaffected. The FPD response for injections ranging from 0.015 to 3 ng was linear with r > 0.999 for all five dialkyl phosphates tested. The minimum detectable level per injection of DEP was 30 pg, and 15 pg for DMTP, DMDTP, DETP, and DEDTP. The recovery data in Table IV was obtained by analysis of urine samples fortified at several levels. The same blank urine sample was used for all fortifications. The low recovery of DEP was the result of both incomplete retention of the metabolite on the extraction cartridges and reduced methylation efficiency in the urine matrix. Minimum levels of detection of the metabolites in urine was 10 ppb for DEP, and 2 ppb for the remaining four metabolites. 2

Table IV. Recovery of dialkyl phosphates from fortified urine samples

ppb

2 10 40 100 200

% recovery ± % standard deviation (n=3) DETP DEP DMTP DMDTP

a

n.d. 8±1 34 ± 7 31 ± 1 27 ± 4

78 90 106 93 84

± ± ± ± ±

23 5 5 5 6

83 83 113 104 96

± ± ± ± ±

98 131 109 96 103

8 4 6 7 7

± ± ± ± ±

9 14 4 9 3

DEDTP

48 79 92 87 87

± ± ± ± ±

7 4 4 6 5

a. n.d. = none detected

Three urine samples of different concentrations were fortified at levels of 20 ppb to test the variability of metabolite recovery from one urine sample to another in our method (Table V). Creatinine content was used as the measure of urine concentration, with 0.4 mg creatinine per ml urine indicating a sample that is quite dilute, and 3.4 mg/ml one that is quite concentrated. These values encompass the range of urine concentrations that might be encountered in a typical study. Average metabolite recovery was similar to that shown in Table IV, with little difference in standard deviation, indicating no effect of urine concentration. Table V. Variation of recovery with urine concentration

creatinine (mg/ml urine)

DEP

0.4 1.1 3.4 avg (%) std. dev. (%)

23 43 25 30 11

% recovery (fortified at 20 ppb) DMTP DMDTP DETP

102 113 104 106 6

95 86 75 85 10

DEDTP

113 97 106 105 8


91 85 86 87 3

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213

The performance of this method on authentic urine samples was examined by analyzing a group of 143 urine samples that were made available from a larger study on farm worker exposure. Concentrations of DMTP, DMDTP, DETP, and DEDTP were determined. Average metabolite concentrations for all compounds were less than 35 ppb (Table VI). Only 16% of the samples tested had no detectable metabolites. This large set of samples did not have a detrimental effect on column or instrument performance. Retention times over the course of the study varied by no more than 3%, and peak areas for standard injections by less than 15%. There was no sign of a general decrease in column, instrument, or detector performance, a problem in some previous procedures (11). Table VI. Results of the analysis of 143 human urine samples

avg (ppb) range (ppb) % positive

DMTP

DMDTP

DETP

DEDTP

6 2 - 27 55%

4 2 - 14 56%

7 2 - 31 41%

3 2-4 6%

Recoveries and limits of detection using the procedure presented here compares favorably with existing methods for DMTP, DETP, DMDTP, and DEDTP. One commonly used method, that of Shafik et al. (6), can detect the 6 common metabolites at levels above 5 ppb. Recoveries from urine fortified at 100 ppb were greater than 95% for all 6 metabolites with this technique. However, several researchers (1.4.5.11) reported poor recoveries and excessive variation in results, indicating that the method may be dependent on the familiarity of the operator with the procedure. Another potential drawback is that only 25-30 samples can be analyzed per week, although this was a significant improvement over existing procedures at the time when this method was developed. The method of Lores and Bradway (8) detects DMP, DEP, DMTP, and DETP at concentrations in urine greater than 10 ppb, and 8-10 samples can be analyzed per day. Recoveries ranged from 40 to 100% when urine was fortified at 500 ppb. Variation in recovery of metabolites from one urine sample to the next averaged 15%. All 6 common dialkyl phosphates may be detected by the method of Reed and Watts (5). The average minimum level of detection in urine is 60 ppb, with metabolite recoveries greater than 90% at fortification levels of 800 ppb. The use of the method described here can result in more rapid analysis of samples compared to existing methods. The time necessary to prepare a single sample for G C analysis is about 15 minutes. Ten samples, the number of cartridges that can be held by our vacuum manifold, can be prepared in forty minutes. Using duplicate standard and sample injections and a three point standard curve, 12 samples can be chromatographed per day. This method is ideally suited to the use of a G C autoinjector, since the chromatographic step is the most time consuming. A single analyst using an autoinjector could easily complete 30-40 samples per day. Existing methods for the analysis of dialkyl phosphates have generally proven inadequate due to long sample preparation and analysis time, poor or variable metabolite recoveries, or lack of sensitivity. Our method is free of these defects in the analysis of DMTP, DETP, DMDTP, and DEDTP. Although DEP recoveries were relatively low, they were consistent enough to yield valid results at urine concentrations over 10 ppb. This method is unsuitable for the analysis of DMP.


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We are seeking to extend this method to the analysis of DMP, and to improve DEP analysis by increasing both the retention of DEP on the extraction cartridges and the derivatization yield. An injector block propylating reagent that produces a unique derivative for inorganic phosphate and each metabolite is being tested and appears promising. Dimethyl propyl phosphate produced upon derivatization of DMP is easily distinguished from the tripropyl derivative of inorganic phosphate. Continuing work on salt addition, pH effects, and solvents for cartridge washing and sample elution should result in better retention of metabolites on the extraction cartridges and cleaner samples. Both effects can improve the efficiency of the method. In summary, we have developed a method for the analysis of urinary dialkyl phosphates that is both rapid and simple. It works well for the four common sulfur containing dialkyl phosphates, and adequately for DEP. DMP cannot be analyzed by this method in its present form. Preliminary work indicates that remaining problems can be overcome without a significant increase in extraction or analysis time.

References 1. Morgan, D. P.; Hetzler, H. L.; Slach, E. F.; Lin, L. I. Arch. Environ. Contam. Toxicol. 1977, 6, 159-173. 2. Franklin, C. A.; Fenske, R. A.; Greenhalgh, R.; Mathieu, L.; Denley, H. V.; Leffingwell, J. T.; Spear, R. C. J. Toxicol. Environ. Health 1981, 7, 715731. 3. Bradway, D. E.; Shafik, T. M.; Lores, E. M. J. Agric. Food Chem. 1977, 25, 1353-1358. 4. Bradway, D. E.; Moseman, R.; May, R. Bull. Environ. Contam. Toxicol. 1981, 26, 520-523. 5. Reid, S. J.; Watts, R. R. J. Anal. Toxicol. 1981, 5, 126-132. 6. Shafik, T.; Bradway, D. E.; Enos, H. F.; Yobs, A. R. J. Agric. Food Chem. 1973, 21, 625-629. 7. Blair, D.; Roderick, H. R. J. Agric. Food Chem. 1976, 24, 1221-1223. 8. Lores, E. M.; Bradway, D. E. J. Agric. Food Chem. 1977, 25, 75-79. 9. Dale, W. E.; Miles, J. W.; Churchill, F. C. J. AOAC 1976, 59, 1088-1093. 10. Miles, J. W.; Dale, W. E. J Agric. Food Chem. 1978, 26, 480-482. 11. Moody, R. P.; Franklin, C. A.; Riedel, D.; Muir, N. I.; Greenhalgh, R.; Hladka, A. J Agric. Food Chem. 1985, 33, 464-467. RECEIVED April 19, 1988


Biological Monitoring for Pesticide Exposure - American Chemical

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