Environ. Sci. Technol. 1982, 16, 250-254
therefore other modes of detection may be subject to interference. Literature Cited Dunn, B. P. In “Polynuclear Aromatic Hydrocarbons: Chemistry and Biological Effects”; Bjorseth, Alf, Dennis, A. J., Eds.; Battelle Press: Columbus,OH, 1979; p 367-377. Brown, R. A,; Weiss, F. T. “Fate and Effects of Polynuclear Aromatic Hydrocarbons in the Aquatic Environment”, Publication No. 4297; American Petroleum Institute: Washington, D.C., 1978. Neff, J. M. “Polycyclic Aromatic Hydrocarbons in the Aquatic Environment: Sources, Fates and Biological Effects”;Applied Sciences: London, 1979. Black, J. J.; Hart, T. F.; Evans, E. In “ChemicalAnalysis and Biological Fats: Polynuclear Aromatic Hydrocarbons“; Cooke, M., Dennis, A. J., Eds.; Battelle Press: Columbus, OH, 1980; p 343-355. Black, J. J. Arch. Environ. Contam. Toxicol., in press. Lopez-Avila,V.;Hites, R. A. Environ. Sci. Technol. 1980, 14, 1382-1390.
Hites, R. A.; LaFlamme, R. E.; Windsor, J. G. Environ. Sci. Res. 1980,16, 397-403.
Grimmer, G.; Bohnke, H. Cancer Lett. (Shannon, Irel.)
Neoplasia”;Kraybill, H. F.; Dawe, C. J., Harshbarger, J. C., Tardiff, R. G., Eds.; N.Y. Academy of Sciences: New York, 1977; p 505-521. Pedersen, M. G.; Hershberger, W. K.; Juchau, M. R. Bull. Environ. Contam. Toxicol. 1974,12, 481-486. Walton, D. G.;Penrose, W. R.; Greene, J. M. J . Fish. Res. Board Can. 1972, 35, 1547-1552. Dressler, M. J . Chromatogr. 1979, 165, 167-206. Benoit, F. M.; LeBel, G. L.; Williams, D. T. Bull. Environ. Contam. Toxicol. 1979,23, 774-778. Alben, K. Environ. Sci. Technol. 1980, 14, 468-470. Saxena, J.; Kozuchowski, J.; Basu, D. K. Environ. Sci. Technol. 1977,11,682-685. Basu, D. K.; Saxena, J. Environ. Sci. Technol. 1978, 12, 795-798. Black, J. J. Roswell Park Memorial Institute, 1980, un-
published data. Department of Environmental Conservation Report on Industrial ChemicalSurvey,Bureau of Industrial Programs; New York State, 1979. Ogan, K.; Katz, E. J . Chromatogr. 1980, 188, 115-127. Glazer, J.; Riggen, R.; Cole, T. In “ChemicalAnalysis and Biological Fate: Polynuclear Aromatic Hydrocarbons”; Cooke, M., Dennis, A. J., Eds.; Battelle Press: Columbus, OH, 1980; p 439-454.
1976,1,75-84.
Leversee, G. J.; Giesy, J. P.; Landum, P. F.; Gerould, S.; Bowling,J. W.; Fannin, T.; Haddock, J.; Bartell, S. Arch. Environ. Contam. Toxicol., in press. Malins, D. C. In “Aquatic Pollutants and Biological Effects with Emphasis on Neoplasia”; Kraybill, H. F., Dawe, C. J., Harshbarger, J. C., Tardiff, R. G., Eds.; N.Y. Academy of Sciences: New York, 1977; p 482-496. Bend, J. R.; James, M. 0.; Dansette, P. M. In “Aquatic Pollutants and Biological Effects with Emphasis on
Received for review June 30,1981. Revised manuscript received November 2,1981. Accepted January 19,1982. This study was supported by a grant from the National Science Foundation and an EPA contract ((2164851 to J.J.B.). Although the research described in this article has been funded wholly or in part by the U.S. Environmental Protection Agency it has not been subjected to the Agency’s required peer and policy review and therefore does not necessarily reflect the views of the Agency, and no official endorsement should be inferred.
Trace Enrichment of Fluorinated Organic Acids Used as Ground-Water Tracers by Liquid Chromatography Klaus J. Stetzenbach,* Stephen L. Jensen, and Glenn M. Thompson Department of Hydrology and Water Resources, University of Arizona, Tucson, Arizona 8572 1, and Laramie Energy Technology Center, PO Box 3395 University Station, Laramie, Wyoming 82071
rn A method was developed for trace enrichment of fluorinated organic acids used as ground-water tracers to allow detection at the low ppb level. By replacement of the injection-valve sample loop with a 3-cm column packed with an octadecylsilane bonded phase, tracer can be extracted from a large water sample as it is pumped through the injection valve. The method increases sensitivity by 3 orders of magnitude while retaining the ease and precision typical of HPLC analysis. Introduction The need to trace ground-water flow and monitor waste-disposal sites has prompted the investigation of several classes of compounds to be used for this purpose (I). This research has resulted in the adoption of a group of fluorinated aromatic organic acids for use as tracer compounds. These compounds are particularly well suited for this purpose because they are exotic in the environment, extremely stable, and not sorbed by the aquifer To whom correspondence should be addressed at Laramie Energy Technology Center. 250
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materials, and they can be separated from other groundwater constituents by bonded-phase high-performance liquid chromatography (BP HPLC). As anions they are readily soluble in water and nonvolatile, which greatly increases the reliability of the quantitative analysis during the monitoring of breakthrough curves. In order to be usable as tracers it is also necessary that these compounds be routinely and accurately measured in the low ppb level or below. The fluorinated aromatic organic acids to be used as tracers are most readily analyzed by means of BP HPLC using a mobile phase of phosphate buffer and methanol, which allows UV detection at wavelengths as low as 200 nm. For this work the absorption bands between 200 and 230 nm provide the best sensitivity. At these wavelengths, standard HPLC methods that permit the direct injection of only 20-200-pL samples typically provide good measurement capability only in the 0.1-1.0 ppm range. Therefore, so that the desired detection limits can be achieved, an on-line precolumn concentrating technique is needed to increase the sensitivity while maintaining the ease and precision typical of HPLC analysis. The removal of organic compounds from large volumes of water by adsorption onto resins followed by elution with
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small volumes of organic solvent has been described by several investigators (2-7). This concentrated sample is then injected onto a GC or LC column for identification and quantification. These methods require a significant amount of sample handling and are also subject to errors stemming from inefficiencies in the concentrating and recovery steps. Bonded octadecylsilane (ODS) and octylsilane (OS) stationary phases have been used to remove organic compounds from water to provide ultrapure water (8,9).Since water is the weakest solvent that can be used on a bonded OS or ODS stationary phase, any dissolved nonionic organic compound that has a low solubility in water will partition strongly into the stationary phase and will not migrate through the column. Long-chained aliphatic and aromatic acids generally have very low solubility in water when they are in the protonated form. In this form they interact with the hydrocarbon bonded stationary phases and will not migrate through the column with a pure aqueous mobile phase (10). Therefore, by use of aqueous solutions buffered to at least 1.3 pH units below the pK, (11) of the dissolved acids, these compounds can be concentrated on a hydrocarbon bonded stationary phase. Preconcentration of water samples directly on the head of standard 25-cm-long analytical columns packed with ODS bonded phases has been reported (12). While this method is appealing because there is little sample preparation and it is extremely simple in principle, the pressure drop across such a column is high, limiting the flow rate that can be used to pump the sample through the column, thus increasing the sample analysis time. More important, however, is the fact that the column is not in equilibrium with the analytical mobile phase at the beginning of the chromatographic run since the sample solvent and the analytical mobile phase cannot have the same composition. This leads to both poor reproducibility of peak-retention times and skewed peaks. This paper reports the results of the use of short (3-4 cm) ODS-packed columns used for preconcentration of ground-water tracers. Similar precolumns have been used for on-line sample enrichment (13),but these techniques require some modifications to the standard HPLC configuration, including the addition of an extra valve. Described herein is a method that can be easily adapted to most HPLC systems as these short columns replace the sample loop on the injection valve allowing the analytical mobile phase to be pumped through the analytical column during the concentration step and keeping the total injected volume independent of the sample volume. The injection volume is always equal to the dead volume of the concentrator column, which is approximately 200-300 pL. Experimental Section Both totally porous 10-pm particle size (RP-18 Merck LiChrosorb, EM Laboratories, Darmstadt, Germany) and pellicular 35-pm particle size (Co-Pell, Whatman Inc., Clifton, NJ) ODS bonded phases were used to pack concentrator columns. The pellicular particles were dry packed into 4 cm X 4.6 mm i.d. 316 stainless-steel columns. The RP-18 packing material was slurry packed into 3 cm X 4.6 mm i.d. 316 stainless-steel columns. In addition, prepacked RP-18 guard columns provided by Rheodyne Inc. (Cotati, CA) were also used as concentrator columns. The concentrating columns were connected to the injection valves in place of the sample loop. Three types of injection valves were used (1)Altex Model 210 (Beckman Instrument Co., Irvine, CAI; (2) Rheodyne Model 7125; (3) Valco Model CV-6-UHPa-N6O (Houston, TX). When the valves were used with a sample loop, they were loaded with the
sample in one direction. When the valve was switched to the inject position, the sample was swept from the loop by the mobile phase in the opposite direction. If the valves are used in this manner with the concentrating columns, then the Altex 210 and the Rheodyne 7125 valves will require special fittings to connect the concentrating pump to the valve load port. This procedure also resulted in a bidirectional flow through the concentrator column. If the concentrating pump is connected to the waste port on the injection valve, no special fittings are required for any of the valves and the flow of the sample solvent and analytical mobile phase through the concentrator column will be unidirectional. The effects of flow direction through the concentrator column are discussed in the Results. Instrumentation All experiments were performed with an Altex dual pump (llOA) gradient-elution liquid chromatograph. Detection was accomplished with a Hitachi Model 155-40 variable-wavelength UV-absorption detector (Beckman Instrument Co., Irvine, CA) set at 200 nm. The data were recorded on a Spectra-Physics Model 4100 computing integrator (Santa Clara, CA). The analytical column was 25 cm long X 4.6 mm id., packed with LiChrosorb RP-18. The concentrating pump was an Altex Model llOA. Reagents Water used for the standards and mobile-phase buffers was prepared by distilling tap water on a Corning Ultra Pure still (Corning, NY). This distilled water was then redistilled over KMn04. Reagent grade methanol was used (Fisher Scientific Co., Fair Lawn, NJ) and was filtered through a Fluoropore 0.5-pm filter (Millipore Corp., El Paso, TX). The potassium phosphate was ACS reagent grade (Mallinckrodt Inc., Paris, KY), and the pentafluorobenzoic acid (PFB) and the m-(trifluoromethy1)benzoic acid (m-TFMBA) were obtained from PCR Scientific (Gainesville, FL). Procedure In the on-line preconcentration configuration, the analytical mobile phase flowed continuously through the analytical column so that the analytical column remained constantly in equilibrium with the mobile phase. With the sampling valve in the load position, the concentrating pump was used to force the sample through the concentrating column. After a predetermined amount of sample was passed through the concentrator column, the concentrating pump was turned off and the injection valve was switched from the load to the inject position. In this position the analytical mobile phase passed through the concentrator column eluting the concentrated material onto the analytical column. Whenever a new sample was used, the concentrating pump was disconnected from the system at the load port of the injection valve and 5-10 mL of the new sample was used to purge the concentrating pump to ensure that no carryover occurred between samples. All internal portions of the sampling valve that carried the sample were flushed by the clean analytical mobile phase during sample injection; therefore, no special valve-cleaning procedure was needed. The concentrating volume was determined by collecting and measuring the effluent from the concentrating column or by timing a given flow rate over a specific time interval. Sample preparation consisted only of filtering the sample through a 0.45-pm filter and buffering the solution to a pH that Environ. Sci. Technol., Vol. 16, No. 5, 1982
251
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Table I. Effect of Concentration Volume on Solute Retention (k' ) Using the Unidirectional-Flow Con figuration m-TFMBA m-TFMBA concd: mL k' concd: mL k' 50 100 PFB concd, mL
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25 pprn m - TFMBA
ANALYTICAL COLUMN
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2OPL o f 25 ppm m - TFM6A CONCENT@ATOR COLUMN And ANALYTICAL COLUMN
49
Figure 1. Comparlson of analytical efficiency between (A) standard Injection of a 20-pL sample directly onto analytical column and (B) analysis using a concentrator column In the unidirectional-flow conflguration.
is 1.3 pH units below the pK, of the compound of interest. The pH adjustment was made with concentrated phosphoric acid so that there was no significant change in sample volume. Results In order to retain the ease and precision associated with HPLC analysis, a precolumn concentration technique should not have any major effects on peak shape (efficiency), solute retention time must be reproducible, and the concentrating capacity must be large enough to increase the sensitivity by several orders of magnitude. These parameters are influenced by the type of stationary phase used in the concentrator column as well as the direction of flow of the sample through the concentrator column. The system configured for unidirectional flow through the concentrator column proved to be superior to the bidirectional-flow configuration. In the unidirectional-flow configuration the trapped solute is eluted from the concentrator column by the mobile phase flowing in the same direction through the column as the water sample. The unidirectional flow method is preferred because it does not disturb the packing bed of the concentrator column. With bidirectional flow, the packing material is shifted during each analysis, resulting in gaps in the packing after 15-20 measurements. Using unidirectional flow, several hundred analyses were performed without any adverse effects. (No experiments have been performed by using the Rheodyne precolumn with bidirectional flow.) Although some adverse effects are associated with the use of bidirectional flow, this method gives greater analytical efficiency for solutes that do not migrate into the precolumn during concentration. With unidirectional flow, the broadening can be minimized by using short, narrow-bore tubing connections and concentrator columns packed with efficient 10-pm particle diameter packing material. Figure 1 shows a comparison of the analytical efficiency of a standard analysis performed by using a 20-pL sample loop with that of an analysis using a concentrator column in the unidirectional-flow configuration. The theoretical plate count of the standard analysis is 25% greater. However, the 25% loss in efficiency is negligible when compared to the increased sensitivity afforded by the concentrating method. Although resolution is also decreased in the concentrating mode, interfering peaks could be removed by adjustments in the mobile-phase parameters. 252
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2.0 2.0
500
k'
PF B concd, mL
1.9
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10 2.6 40 2.1 25 2.2 45 2.1 30 2.2 50 2.1 35 2.1 a For rn-(trifluoromethy1)benzoicacid (pK, = 3.9). For pentafluorobenzoic acid (pK, = 1.74).
The 10-pm RP-18 packing material proved to be far superior to the pellicular in both capacity and efficiency. With the unidirectional-flow configuration, the sample is also chromatographed on the concentrator column, so packing both the concentrator and analytical columns with the same packing materials ensures uniform elution behavior. The pressure drop across these short columns is not very high. With concentrating flows of 10 mL/min of water, the pressure drop is approximately 1000 psi. Flow rates as high as 25 mL/min were employed during the concentrating procedure by using the Altex llOA pump equipped with a prep head. During the elution and analysis step using a mobile-phase composition of 50/50 methanol/water at a flow rate of 2 mL/min, the concentrator column adds an additional 400-500 psi of pressure to the system. The RP-18 material proved to be the preferred packing for the concentrator columns due to the greater soluteretention capacity of this material. This packing enabled rn-TFMI3A (pK, = 3.9) to be quantitatively extracted from sample volumes as large as 500 mL. Much larger volumes are possible if time for concentration is not a limiting factor. The concentrator columns packed with the pellicular packing material did not have the capacity necessary to achieve the desired sample enrichment. A weak acid (pK, = 3.9) started to elute from this packing material with concentrating volumes of 25 mL, and a strong acid (pK, = 1.74) was not retained. Experiments were performed with different concentrating volumes to determine the effect of sample size on solute-retention time by using the unidirectional-flow configuration. If the solute migrates through the concentrating column, the retention time (K )' will decrease with increasing sample size. Since the retention time for rn-TFMBA is nearly constant (Table Ia) for all concentrating volumes, it can be assumed that this acid is concentrated at the head of the concentrator column. However, with a stronger acid such as pentafluorobenzoic acid (PFB) (pKa = 1.74), breakthrough occurred after 40 mL of sample had been passed through the concentrator column. The stronger acid migrates through the concentrator column during the concentrating procedure, as evidenced by the fact that the retention time decreased as the sample size increases (Table Ib). These experimental observations are consistent with the theories of acid-base chemistry and the retention mechanisms of hydrocarbon bonded stationary phases (10, 14,15). For retention of acids on ODS bonded phases with an aqueous mobile phase, they cannot be in the ionic form and they must be water insoluble in their nonionized form. Therefore, the sample pH must be at least 1.3 units below the pKa of the
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solute. The pH of the sample used in these experiments was 2.2-2.5, which is more than 1.3 units below the pK, of rn-TFMBA (pK, = 3.9) but higher than the pK, of PFB (pK, = 1.74). The sample pH could be dropped below pH 2 when the need arises to concentrate stronger acids, but this low pH will have a strong detrimental effect on the column life. While this limits the use of very large concentrating volumes to acids that have pK, values of 3.3 or larger, some concentrating can be achieved with stronger acids. Figure 2 shows the peak-area response to various concentrating volumes of PFB. This calibration curve is linear and usable up to concentrating volumes of 40 mL. Volumes larger than 40 mL exceed the capacity of the concentrator column, resulting in loss of sample. Longer concentrator columns would increase the amount of sample that can be concentrated. A calibration curve is normally developed by concentrating fixed volumes of two or three standard solutions as shown in Figure 3. However, with this concentrating method it is also possible to use one standard and vary the concentrating volumes when creating a calibration curve. If the tracer is quantitatively trapped on the concentrator column, the detector response will be a function of the mass of solute injected only and will be independent of the amount of concentration of the sample. To verify that the system was providing quantitative trapping and recovery, the calibrations curve (Figure 4) was constructed with samples of 5-50 mL of a 25-ppb rn-TFMBA standard solution. The extraction efficiency was checked by comparing the peak-height response of 250- and 550-mL concentrating volumes of a 0.5-ppb solution to the calibration curve obtained with the 25-ppb standard. The 25-ppb solution is 50 times as concentrated as the 0.5-ppb solution. The 250-mL concentrating volume of the 0.5-ppb solution produced the same response as 5 mL of the 25-ppb solution. A 550-mL sample of the 0.5-ppb solution gave a response equivalent to 11 mL of the 25ppb standard. In each case where the average response from triplicate
GRGUM WATER SAMPLE f 8 70 ppbJ 2 288 Y
Figwe 5. Chromatogram of a wound-water S a w c~htdkihgm TFMBA. A sample was armbed on a W-18 andykal u h m wkh 40% methanol and 60% 0.01 M phssphete CJ&& @H 7.1) 81 2 W m i n . Detection was accompRshed with a vadabb W detector set at 200 nm.
analysis of a given mass of solute injected frahr the d.5-pOb solution was compared to the same mass iuljected f W ithe 25-ppb solution, the difference or error w89 iesS thdn 4 % .
Conclusion This method of sample enrichment has b&n Used for analysis of rn-TFMBA, PFB, and benzoate, which dCre used as ground-water tracers for several tracing teeta at two different sites. The enrichment procedure proved tb be rapid, simple, and efficient, and no interfereh- were found in these natural ground waters. Figure 5 ehodts a chromatogram of a 10-mL sample of ground waur that contained 8.7 ppb of rn-TFMBA. The reltltive stahdard deviation for this method on over 30 me&ufemetlfs of ground-water samples analyzed in this laboratory is &3% (1 a).
While only organic acids were used in this emdy, it is believed that this enrichment technique is a p p h b l e to a wide range of compounds that can be ertracted from water into a hydrocarbon phase. For cbhpollnds thbt partition strongly into the hydrocarbon phase (such 88 the weak acids), very large concentrating volurtieu, which wult in increased sensitivity of 3 or more orders of m w i t u d e , can be used. Acknowledgments
We express our appreciation to Den& BkMas for packing many concentrating and analytical columns and the Steve Bakalyar for generously providing the Rhcodyne Environ. Sci. Technol., V d . 16, No. 5, 1g82
253
Environ. Sci. Techno/. 1982, 16, 254-262
precolumns. This work was supported by the U.S. Nuclear Regulatory Commission under Contract No. NRC-04-78275. Literature Cited (1) Davis, S.N.; Thompson, G. M.; Bentley, H. W.; Stiles, G. Ground Water 1980,18,14-23. (2) Thurman,E. M.; Malcolm, R. L.; Aiken, G. R. Anal. Chem. 1978,50,775-779. (3) Aiken, G. R.;Thurman, E. M.; Malcolm, R. L. Anal. Chem. 1979,51,1799-1803. (4) Koch, D.L.;Kissinger, P. T. Anal. Chem. 1980,52,27-29. (5) Creed, C. G.Res.lDev. 1976,279,40-44. (6) Cassidy, R.M.; Elchuk, S. J. Chromatogr. Sci. 1980,18, 217-223. (7) Saner, W. A.; Jadamec, R. T.; Sager, R. W. Anal. Chem. 1979,51,2180-2188.
(8) Simpson, R. L. J. Am. Lab. (Fairfield,Conn.) 1977,9,5, 109-115. (9) Gurkin, M.; Ripphann, J. Am. Lab. (Fairfield,Conn.) 1980, 12,5, 99-102. (10) Horvath, C.; Melander, W.; Molnar, I. Anal. Chem. 1977, 49,142-154. (11) Freiser, H.; Fernando, Q.“Ionic Equilibria in Analytical Chemistry”;Wiley: New York, 1963;Chapter 4. (12) Kirkland, J. J. Analyst (London) 1974,99,859. (13) Van Vliet, H. P. M.; Bootaman, Th. C.; Frei, R. W.; Brinkman, U. A. J. Chromatogr. 1979,185,483-495. (14) Pietrzyk, D. J.; Chu, C.-H. Anal. Chem. 1977,49,860-867. (15) Deming, S.N.; Turoff, M. Anal. Chem. 1978,50,546-548.
Received for review May 6,1981. Revised manuscript received December 21,1981. Accepted December 21,1981. Thk work was supported by the U S . Nuclear Regulatory Commission under Contract No. NRC-04-78-275.
Formaldehyde and Other Carbonyls in Los Angeles Ambient Air Daniel Grosjean
Environmental Research & Technology, Inc., 2625 Townsgate Road, Westlake Village, California 9 1361 From selective sampling and liquid chramatography analysis, ambient levels of carbonyl compounds as 2,4dinitrophenylhydrazoneshave been measured in the Los Angeles area during severe photochemical pollution episodes. Gas-phase concentrations and diurnal profiles are presented for six carbonyls: formaldehyde (up to 48 ppb), acetaldehyde (135ppb), propanal (114 ppb), butanal (17 ppb), 2-butanone (114 ppb), and benzaldehyde (11ppb). Also presented are particulate-phase concentrations and particle/gas distribution ratios for five carbonyls. Ambient carbonyl levels are discussed with respect to anthropogenic emissions and to photochemical production and removal in polluted air. Advantages and current limitations of the method employed are briefly discussed. H
Introduction
Of the major classes of organic compounds involved in photochemical air pollution, carbonyls (aldehydes and ketones) are of critical importance as products of the photooxidation of gas-phase hydrocarbons, as a major source of free radicals, and as precursors to organic-aerosol formation in urban air. Over the past several decades, considerable progress has been achieved, through both theoretical and experimental studies, in elucidating the chemical pathways involved in the formation and removal of carbonyl compounds in the atmosphere (1). The importance of carbonyls is reflected, for example, in the fact that detailed computer kinetic chemical models employed in oxidant (ozone) formation simulations and other regulatory applications all require formaldehyde and other carbonyls as data input (2). However, due to a large extent to analytical difficulties, measurements of ambient levels of carbonyl compounds have been limited to a few urban areas, and in most cases only formaldehyde and/or “total aldehydes” measurements have been performed. Using a method recently developed in our laboratory (3, 4 ) , we have performed measurements of ambient levels of carbonyl compounds in the Los Angeles area, under conditions of moderate to severe photochemical pollution, including the worst smog episodes encountered in 1980 in the Los Angeles air basin. 264
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Methods
The method employed in this study (3,4)entails the use of a selective reagent 2,4-dinitraphenylhydrazine(DNPH), which reacts with carbonyl compounds to form 2,4-dinitrophenylhydrazones (hereafter referred to as hydrazones) according to the reaction shown in Figure 1. After organic solvent extraction, the hydrazones are separated by high-pressure liquid chromatography (HPLC) and quantitated on the basis of their absorption at 360 nm. Quantitative aspects of the method including analytical recovery, sampling efficiency, reproducibility, and detection limits have been previously reported ( 3 , 4 )and only a brief description of the sampling and analytical protocols employed in this study is presented here. Samples were collected with microimpingers containing 10 mL of an aqueous, acidic (2 N HC1) solution of DNPH and 10 mL of a 9:l by volume mixture of cyclohexane and isooctane. In the absence of organic solvents, quantitative recovery is obtained for formaldehyde but not for other aliphatic and aromatic carbonyls (5). With organic solvents added, recoveries for these carbonyls are essentially complete and result from in situ extraction of the hydrazone in the organic phase, whose effect is to displace the aqueous-phase equilibrium (Figure 1)toward hydrazone formation. Detailed collection efficiency studies for formaldehyde, acetaldehyde, and benzaldehyde in the range of concentrations (a few micrograms) relevant to ambient-air sampling are described elsewhere (3,5). On several occasions, including nighttime sampling, an automated sampling device was employed as previously described (6). Typically, 60 L of ambient air was collected at a measured flow rate of -1 L mi&. Blanks were included with each batch of DNPH reagent and shipped to and from the field along with actual samples. Also included were control samples spiked with known amounts of acetaldehyde 2,4-dinitrophenylhydrazone. Samples, blanks, and controls were returned to the laboratory at low temperature in the dark and were stored at refrigerator temperature prior to extraction and HPLC analysis. After addition of an internal standard, extraction with a mixture of hexane and methylene chloride, and solvent
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