Evaluation of Media and Derivatization Chemistry for Six Aldehydes in

Environ. Sci. Technol. 2001, 35, 2301-2308

Evaluation of Media and Derivatization Chemistry for Six Aldehydes in a Passive Sampler L.-J. SALLY LIU,* RUSSELL L. DILLS, MIKE PAULSEN, AND DAVID A. KALMAN Department of Environmental Health, University of Washington, Box 357234, Seattle, Washington 98195-7234

We evaluated the GMD passive sampler for its suitability to measure six aldehydes over a 7-d period in population exposure studies. The six target aldehydes were formaldehyde, acetaldehyde, acrolein, crotonaldehyde, glyoxal, and methylglyoxal. The GMD sampler contains a silica gelimpregnated cellulose pad coated with 2,4-dinitrophenylhydrazine (DNPH) hydrochloride. This agent reacts with formaldehyde to form a hydrazone that is quantified with a high-performance liquid chromatograph. The GMD sampler was tested for background contamination and aldehyde recoveries after 0, 1, and 7 d of storage. Results indicated that the GMD monitor, as currently manufactured, is suitable for shorter-term sampling (up to 24 h) of formaldehyde and acetaldehyde. It is however not acceptable for sampling of acetaldehyde, acrolein, crotonaldehyde, glyoxal, and methylglyoxal over a 7-d exposure period due to the chemical reactions on the silica gel-impregnated cellulose pad. Glyoxal- and methylglyoxal-DNPH derivatives formed on the cellulose and Teflon-coated glass fiber pads that had been prepared with glycerol under acidic and oxidative conditions. Acrolein- and crotonaldehydeDNPH derivatives diminish through the reverse reaction of the DNPH derivatives to form free aldehydes under acidic conditions. We showed that the unknown reaction products of acrolein and crotonaldehyde derivatives were not pyrazolines but probably resulted from E/Z isomerization. These conversion reactions are favored in acidic conditions present in either the derivatization solution or the collection medium. The most consistent recovery was obtained on glass fiber pads. In particular, recoveries of crotonaldehyde- and acrolein-DNPH derivatives were increased through the use of a pH 4 buffered derivatization solution. These chemical instability problems were overcome by using a pH 4 buffer (citric acid/sodium citrate) and an alternative hygroscopic agent (1,3-butanediol) in the DNPH derivatization solution. Results with DNPH derivatives from these spiking experiments were further confirmed with gas-phase spiking experiments. We determined the optimal acidity, buffer solution, and concentrations of the buffer solution and 1,3-butanediol for the DNPH derivatization solution. This new formulation of the DNPH derivatization solution can be used for collection of the six target aldehydes over a 7-d sampling period.

Introduction Aldehydes in the environment result from natural and industrial processes. With the use of alternative fuels and additives derived from methanol and ethanol, ambient exposures to aldehydes from automotive sources are expected 10.1021/es001795c CCC: $20.00 Published on Web 04/19/2001

 2001 American Chemical Society

to increase (1). Several aldehydes are targeted as potentially important toxicants by the Clean Air Act Amendments of 1990. However, little quantitative information on personal exposures to aldehydes exists for forming reliable estimates of human health risks from aldehyde exposures. Very few studies have been conducted in the United States to characterize aldehyde exposures and health effects in the general population. One of the major limiting factors is the lack of an adequate long-term (1 week or longer) personal sampler that can monitor exposures to several aldehydes at low cost. Passive sampling devices for carbonyls generally rely on a substrate coated with a reagent to chemically trap the analytes as nonvolatile derivatives. Derivatization reagents include sodium hydrogen bisulfite, 2,4-dinitrophenylhydrazine (DNPH), and chromotropic acid. Conventional passive sampling devices generally consist of a thin, filter-like collection element inside a housing with a diffusive barrier. Of those reviewed, all (with one exception) have only been validated for formaldehyde. These include the GMD badge (Bacharach Inc., Pittsburgh, PA), 3M badge 3721 (3M, Inc., Minneapolis, MN), SKC badge (526 series) (SKC, Inc., Fullerton, CA), Ferm passive sampler (2), and ChemDisk badge (Assay Technology, Palo Alto, CA). The exception was the last of these, which was also validated for glutaraldehyde. In the past 2 yr, new designs of passive samplers containing a solid adsorbent have been developed to allow for collection of multiple carbonyls. These include the sampling tube for acetaldehyde, acetone, 2-butanone, and cyclohexanone (3); a diffusive sampling tube for formaldehyde, acetaldehyde, and acetone (4); a passive badge for valeraldehyde and acrolein (5); and a syringe-style sampler for formaldehyde, acetaldehyde, acetone, acrolein, propionaldehyde, crotonaldehyde, benzaldehyde, and hexaldehyde (6). Various adsorbents and coating agents have been utilized. The Binding (3) and Uchiyama and Hasegawa (4) samplers contain silica gel coated with 2,4-DNPH; the Tsai and Hee sampler (5) uses a Tenax TA pellet coated with 10% (w/w) O-(2,3,4,5,6pentafluorobenzyl)hydroxylamine hydrochloride; and the Zhang et al. sampler (6) contains C18-silica packing coated with dansylhydrazine. None of these samplers has been tested for chemical stability for various aldehydes for more than 24-h exposure durations, with one exception (Zhang et al. sampler was tested for 48 h). The utility of carbonyl derivatization with 2,4-DNPH for air sampling is well-established. Samplers can be extracted with acetonitrile; extracts are then analyzed using a highperformance liquid chromatograph (HPLC) (3, 7-10) or a liquid chromatograph/mass selective detector (11). However, the stability of easily degraded aldehydes (such as acrolein, crotonaldehyde, glyoxal, and methylglyoxal) or their DNPH derivatives on collection media is unknown. Reports of air sampling with 2,4-DNPH-impregnated media typically used active methods (with pumps) and short duration of less than 24 h (4, 6-10, 12). These studies used either C18-silica or silica collection media that was stored frozen after collection until chemical analysis was completed. To meet the needs of long-term exposure assessment studies in large populations, a stable and convenient personal sampler is necessary. This sampler should be suited for exposure of 7 d or longer and should tolerate being stored or shipped at room temperature. Our study examines the feasibility of using the GMD badge and the DNPH reaction * Corresponding author e-mail: [email protected]; phone: (206)543-2005; fax: (206)543-8123. VOL. 35, NO. 11, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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derivatizing system for sampling of formaldehyde, acetaldehyde, acrolein, crotonaldehyde, glyoxal, and methylglyoxal under long-term (7-d) monitoring conditions.

Experimental Methods Materials. DNPH derivatives of glyoxal and methylglyoxal were synthesized. The identity and purity of these two derivatives were confirmed by melting point, HPLC, and solidprobe mass spectrometry. The dihydrazone derivatives were formed. The cyclized forms of the 2,4-dinitrophenylhydrazone of acrolein and crotonaldehyde were synthesized from the aldehydes, hydrazine, and 2,4-dinitrochlorobenzene (13). Acrolein formed 1-(2,4-dinitrophenyl)-2-pyrazoline and crotonaldehyde formed the 1-(2,4-dinitrophenyl)-5-methyl-2pyrazoline. Identities were confirmed by melting point and HPLC-MS. Sampler. The GMD badge was selected for testing due to its simplicity and desirable features for use in population studies. It has been shown to have high sensitivity at low concentrations of formaldehyde and low wind speeds (14, 15). It is easy to use and handle by subjects. In addition, each sampler comes with separate sampling and blank control compartment. The polypropylene GMD sampler weighs 0.5 oz (60 × 30 × 5 mm3). The sampler contains a silica gelimpregnated cellulose pad (20 × 45 mm2), coated with 2,4DNPH solution containing glycerol, ethanol, DNPH, phosphoric acid, and acetonitrile (15). The pad is placed beneath a 2.9-mm-thick screen. The pad is embossed into two sections by a small ridge on the screen plate. Half of the screen contains 112 holes, and the pad beneath the holes is used for sampling while the other half is never exposed and is used as the blank control. A sliding cover seals the holes when the sampler is not in use. Each unused sampler is provided in a Mylar package with a DNPH-coated scavenger pad. Sample Analysis. Extraction of the 2,4-DNPH derivatives was performed by mechanical agitation of the pad with 3 mL of acetonitrile in a 4-mL glass vial. 2,4-DNPH derivatives were analyzed by an HPLC (HP1050; Avondale, PA) equipped with a variable wavelength detector and an autosampler. We selected the Adsorbosphere UHS (5 µm packing, 4.6 × 150 mm; Alltech, Deerfield, IL) reverse-phase C18 column because of its ability to give excellent baseline resolution of acrolein and acetone. A mobile phase with a tertiary gradient utilizing acetonitrile, methanol, and water provided resolution of target aldehydes and the background contaminants encountered (16). Initial conditions for mobile phase (33% water [containing 3% acetonitrile], 7% acetonitrile, and 60% methanol) were held for 7 min. A linear gradient to 14 min produced a final composition of 10% water [containing 3% acetonitrile], 10% acetonitrile, and 80% methanol. This composition was held to the termination of the run at 17 min. Column temperature was 50 °C. Flow rate was 1 mL/ min. Equilibration time between runs was 5 min. Detector wavelength was initially 360 nm and was switched to 440 nm at 13 min for the detection of the dicarbonyls at their optimum wavelength. Injection volume was 10 µL. Calibration standards of aldehyde-DNPH derivatives for formaldehyde, acetaldehyde, acrolein, and crotonaldehyde were purchased from Supelco (Bellefonte, PA). Quantitation was performed by external standard calibration using a sixpoint curve ranging from 0.05 to 10 ppm. Detector response was linear over this range as determined by R 2 values greater than 0.999 for each analyte. Evaluation of Unmodified GMD Sampling Media. We found significant levels of the target aldehydes and other aldehydes in samplers (N ) 10) at or beyond their expiration date. Subsequently, newly made GMD samplers (N ) 4) were evaluated for background contamination. Samplers were tested for recovery and stability of aldehyde-DNPH derivatives by removing the silica gel-impregnated cellulose pads 2302

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from the sampler, spiking the pad with the target aldehydeDNPH derivatives, and reassembling the sampler. As a control for sampler housing-derived contamination, some pads were placed in glass vials with Teflon-lined caps. The samplers and glass vials were sealed and stored at room temperature. The pads were removed from the samplers and glass vials at three times: day 0 (immediately after spiking), day 1 (after 24 h of storage), and day 7 (after 168 h of storage) and were analyzed for aldehyde-DNPH derivatives. Evaluation of Alternative Hygroscopic Reagents. The increase of glyoxal and methylglyoxal over time was hypothesized to be due to the oxidation of glycerol. Glyoxal, methylglyoxal, and acrolein can be produced from glycerol under acidic and oxidative conditions (17, 18). We tested the possibility that glycerol was the chemical source of glyoxal and methylglyoxal in the DNPH derivatization solution. DNPH solutions were prepared, one with glycerol and the other with 1,3-butanediol, as a replacement for glycerol. Silica gel-impregnated cellulose pads (20 × 22.5 mm) were coated with solutions and spiked with aldehyde-DNPH derivatives. The pads were stored at room temperature and analyzed after 1 and 7 d (N ) 2 for each solution type and each storage duration). Collection Medium Tests. We examined the stability of aldehyde-DNPH derivatives on various collection media. The test media were chosen to be either more inert than the silica gel-impregnated pads or similar to packing in the C18 Sep-Pak cartridges. The media included Teflon-coated glass fiber filters (Fiber Film, T60A20, 7215, Pall Gelman Laboratory, Ann Arbor, MI), cellulose fiber pads (Gelman, 66239), glass fiber filters (type A/E, 1 µm pore, 61638; Pall Gelman Laboratory), quartz microfiber filters (Whatman QM-A, 1851037; Clifton, NJ), C18 silane-treated (octadecyldimethyl chlorosilane) silica gel-impregnated cellulose, Teflon-fiber mat (0.3 mm thickness) (grade PF10X, Polyflon, PF0508X10IN; Advantec MFS, Pleasanton, CA), and C18-silica thin layer chromatographic or TLC plates (C18 silica, 200 µm thick, glass backed, J. T. Baker, VWR, West Chester, PA). For these experiments, up to six pieces of test media (20 × 22.5 mm) were coated with DNPH solutions formulated with either 1,3-butanediol or glycerol. After being spiked with aldehydeDNPH derivative solutions in tetrahydrofuran (80%) and acetonitrile, samples were stored at room temperature and analyzed after 24 h (day 1) and 168 h (day 7). Optimization Tests. After the best performing medium (glass fiber filters) and a better hygroscopic agent (1,3butanediol) were selected, three serial experiments were performed to optimize pH and the concentrations of reagents in the DNPH derivatization solution, with respect to collection efficiency for the target aldehydes. The main purpose was to test the stability of aldehyde-DNPH derivatives. Thus, other than introducing water vapor to eliminate dry air artifacts (19) in two of the experiments, complicated environmental factors such as the well-known ozone artifacts (20) were not introduced to the system. The collection efficiency was determined as the ratio of the aldehyde concentration in acetonitrile extracted from the medium to the aldehyde concentration in spiking solution. The first test evaluated the collection efficiencies of aldehydes in 22 different derivatization solutions. These included three buffers (citric acid/sodium citrate, phosphoric acid/sodium hydroxide, and hydrochloric acid/potassium chloride), five pH levels (pH 1 to pH 5), and four butanediol concentrations (0.4, 1, 5, and 10% v/v). The buffer concentration in the derivatization solutions was 10 mM with the exception of citrate buffer (pH 4), for which additional 20 and 100 mM citrate buffers were included. DNPH concentration was fixed at 5 mg/mL. Three replicate glass fiber pads were prepared for each solution. Pads were washed with acetonitrile, soaked in the derivatization solution for 1 min,

FIGURE 1. Diminishing of acetaldehyde-, acrolein-, and crotonaldehyde-DNPH derivativatives through two possible mechanisms: (a) Cyclization of the alkene hydrazone to form a pyrazoline acts as a potential source for diminishing of acrolein (RdH) and crotonaldehyde (RdCH3). (b) Acid-catalyzed E/Z isomerization accounts for the appearance of new peaks in chromatograms of the asymmetrical hydrazones. and allowed to dry in a nitrogen-purged glovebag. Two identical Tedlar bags were made, and each was loaded with 33 pads. The bags were sealed with a thermal impulse sealer, filled with 20 L of air, and then spiked with liquid-phase aldehydes through a septum, which evaporated spontaneously in the bag. The exposure duration was 24 h for this and the subsequent optimization tests. The calculated result was the aldehyde concentration measured in the pad relative to the aldehyde concentration in the spiking solution. The second test determined the optimal concentrations of butanediol and the best buffer (citric acid/sodium citrate buffer at pH 4). Twenty-four DNPH derivatization solutions were made to test six butanediol (0.1, 1, 5,10, 20, and 50% v/v) and four buffer (1, 10, 20, and 50 mM) concentrations. For each solution, five replicate glass fiber pads were coated. The pads from each solution were evenly distributed in five Tedlar bags. Water was injected into the bags through septa and evaporated spontaneously to generate a defined relative humidity (RH) (∼25%). The third optimization test was to confirm the optimal concentration of butanediol in the DNPH derivatization solution at ∼25% RH. Three levels of butanediol (5, 10, and 20% v/v) were tested in 50 mM citric acid/sodium citrate buffer at pH 4 (the best combination from test 2). Six pads were coated with each solution. Pads were exposed to aldehydes inside a sealed Tedlar bag. Results were shown as the amount of aldehyde-DNPH derivatives in acetonitrile extract from the exposed pads.

Results and Discussion Chemical Anomalies. When acrolein-, crotonaldehyde-, and acetaldehyde-DNPH derivatives were stored for 7 d on strongly acidic silica media, such as in the unmodified GMD sampler, the original hydrazone peak diminished with the corresponding appearance of another peak. Tejada (21) noted, without an explanation, a similar stoichiometric behavior on silica gel cartridges coated with acidified DNPH. For each of these aldehydes, the new peak was indistinguishable from the original by mass spectroscopy. For the unsaturated aldehyde derivatives, cyclization of the alkene hydrazone to form a pyrazoline was a possible explanation for the new peaks (Figure 1a). We synthesized the pyrazolines and found that they had the same mass spectra as the uncyclized hydrazones but eluted several minutes earlier. Acidification of acetonitrile solutions of acrolein-, crotonaldehyde-, or acetaldehyde-DNPH derivative was found to cause a partial transformation of the derivatives matching the new chromatographic peaks (Figure 2). Karabatsos et al. (22) and Tayyari et al. (23) noted the acid-catalyzed isomerization of hydrazones in solution to specific ratios of E to Z

isomers dependent upon the solvent and the compound. We concluded that the most likely explanation for the appearance of new peaks in the chromatogram for the asymmetrical hydrazones was acid-catalyzed E/Z isomerization (Figure 1b). Sampler Tests. New GMD samplers, as routinely manufactured, are contaminated with acetaldehyde, acrolein, glyoxal, and methylglyoxal (Table 1). This prevented us from using them for environmental sampling. To determine the sources of aldehydes, we analyzed individual components of the sampler and its aluminum Mylar package. No detectable aldehydes were found in any sampler components nor on the package, except for the DNPH-coated pads, including the sampling pad inside the sampler body and the “scavenger” pad. Both pads showed similar levels of contamination, although the sampler pad was sealed within the sampler body while the scavenger was exposed within the package. We concluded that the contaminants must have originated from the derivatization solution, the sampler preparation processes, or chemical reactions occurring on the coated pad. Contamination with acrolein and glyoxal was eliminated by the sampler manufacturer through procedural changes. Background contamination of methylglyoxal disappeared when a more purified grade of glycerol was used. The only remaining background contaminant was acetaldehyde, which ranged between 0.10 and 0.15 µg/mL in acetonitrile extracts (3 mL) of unexposed sampler pads. Coated sampler pads with the improved background levels were used for the subsequent recovery and stability tests of the GMD sampler. The recoveries of aldehyde-DNPH derivatives on spiked pads are shown in Figure 3. On day 0, most aldehyde-DNPH derivatives, except for acrolein and crotonaldehyde, showed a near 100% recovery. After 24 h of storage at room temperature, all hydrazones but glyoxal- and methylglyoxalDNPH derivatives exhibited a decline in recovery. After 7 d of storage at room temperature, 0% recoveries were observed for the unsaturated aldehydes (acrolein and crotonaldehyde), while formaldehyde- and acetaldehyde-DNPH derivatives showed recoveries of 47% and 74%, respectively. Glyoxal and methylglyoxal recoveries increased to approximately 190%. The same recovery experiments with similar results were performed on four Assay Technology samplers, which were coated with a similar DNPH solution. After 7 d in storage, acrolein and crotonaldehyde collected on the Assay Technology samplers decreased to 36% and 81% in recovery, respectively, while glyoxal and methylglyoxal increased to a recovery of 304% and 226%, respectively. We hypothesized that the decreases in recoveries of the hydrazones were caused by nonspecific chemical reactions on the collection medium and the acid-catalyzed reverse reaction of the hydrazones. Acid is known to catalyze the reverse reaction (aldehyde-DNPH f aldehyde + DNPH) (24, 25). At low pH (e2), the reverse reaction is favored. Bicking et al. (26) suggested that a pH of 4 is optimum for the formation of the 2,4-dinitophenylhydrazones. In one of our experiments, phosphoric acid equivalent to the amount in the DNPH derivatization solution was added to the aldehyde-DNPH derivatives in an acetonitrile solution. Results showed 20-30% decreased recoveries in aldehydeDNPH derivative after 7 d of storage at room temperature. Collection Media and Hygroscopic Agents. Alternative hypotheses for the instability of some aldehyde-DNPH derivatives were that the inherent acidity or chemical activity of the silica gel-impregnated cellulose pad may either enhance the decomposition rate of DNPH derivatives or irreversibly bind the derivatives. Under these assumptions, treating the pad with various deactivation agents would increase stability of aldehyde-DNPH derivatives on the pad. We soaked the pads in one of the following solutions: VOL. 35, NO. 11, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 2. Aldehyde-DNPH derivative controls and isomers after 7-d incubation in DNPH solution with phosphoric acid: (a) acroleinDNPH derivative, (b) acrolein-DNPH derivative and isomer (peak at retention time 11.357 is acetone-DNPH derivative), (c) crotonaldehydeDNPH derivative, and (d) crotonaldehyde-DNPH derivative and isomer. dimethyldichlorosilane (followed by methanol), triethanolamine, ammonium hydroxide, EDTA, or sodium bicarbon2304

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ate. These treated tapes were then dried in vacuo, coated with DNPH derivatization solution, spiked with aldehyde-

TABLE 1. Background Contamination (Shown in Mean ( Standard Deviation) in Four Randomly Selected New GMD Samplers

formaldehyde acetaldehyde acrolein crotonaldehyde glyoxal methylglyoxal

extract concn (µg/mL)

air concna (µg/m3)

NDb 0.10 ( 0.02 0.09 ( 0.02 ND 0.52 ( 0.04 2.78 ( 0.11

ND 1.46 ( 0.33 1.43 ( 0.24 ND 8.45 ( 0.65 50.8 ( 2.05

a Assuming 7-d sampling at the theoretical collection rate based on the measured collection rate for formaldehyde. The extraction volume is 3 mL. b ND, not detected.

FIGURE 4. Recovery of aldehyde-DNPH derivatives on spiked collection media coated with DNPH solution with glycerol after (a) 1 and (b) 7 d in storage (N ) 2 per pad type).

FIGURE 3. Recovery of aldehyde-DNPH derivatives on spiked GMD samplers after 0, 1, and 7 d in storage at room temperature (N ) 3 per symbol). DNPH derivatives, stored at room temperature, and then analyzed after 1 or 7 d. None of the above treatments improved the recovery of acrolein and crotonaldehyde (day 1 recoveries of 10-40%). Deactivation of the pad by these procedures therefore did not appear to improve stability of derivatives. We then tested various collection media in search of the most inert medium toward DNPH derivatives for aldehyde collection. Figure 4a,b shows aldehyde recoveries on various spiked collection media, including cellulose, quartz, Tefloncoated glass fiber (T-Glass), and glass fiber pads. On spiked cellulose and quartz pads, less than 70% of acrolein and crotonaldehyde derivatives were recovered after 1 d of storage at room temperature (Figure 4a). After 7 d of storage, recoveries from the spiked cellulose pads were less than 50% for acrolein- and crotonaldehyde-DNPH derivatives but more than 160% for glyoxal- and methylglyoxal-DNPH derivatives. These trends were very similar to those observed in Figure 3. Recoveries from the Teflon-coated glass fiber pads showed similar trends for these four aldehydes but with higher recoveries. Quartz pads, which are typically considered more inert than glass fiber pads, had the lowest recoveries of acrolein and crotonaldehyde on day 1, a low recovery of crotonaldehyde on day 7, but high recovery of acrolein on day 7. Glass fiber pads seem to have the most stable recoveries for most aldehyde derivatives, except for crotonaldehyde. The same four media were coated with a DNPH solution containing 1,3-butanediol, rather than glycerol as the hygroscopic agent, spiked with aldehyde-DNPH derivatives,

FIGURE 5. Recovery of aldehyde-DNPH derivatives on spiked collection media coated with DNPH solution with 1,3-butanediol after (a) 1 and (b) 7 d in storage (N ) 2 per pad type). and tested for recoveries (Figure 5a,b). The most notable difference from the experiments with glycerol (Figure 4) was that no dramatic increases in glyoxal and methylglyoxal in cellulose and Teflon-coated glass fiber pads were observed. Glass fiber and Teflon-coated glass fiber pads provided the best recoveries for all aldehydes, especially for acrolein and VOL. 35, NO. 11, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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TABLE 2. Recovery (%) of Aldehyde-DNPH Derivatives on Spiked Glass Fiber Pads Coated with DNPH/1,3-Butanediol and Citric Acid/Sodium Citrate Buffer at pH 4

formaldehyde acetaldehyde acrolein crotonaldehyde glyoxal methylglyoxal

day 1 (N ) 6)

day 7 (N ) 6)

85 ( 10 97 ( 2 100 ( 2 92 ( 4 102 ( 4 93 ( 1

89 ( 4 102 ( 4 93 ( 1 95 ( 1 105 ( 2 95 ( 2

TABLE 3. Top Five Collection Efficiencies for Each Aldehyde from the First Optimization Testa

aldehyde formaldehyde

acetaldehyde

FIGURE 6. Recovery of aldehyde-DNPH derivatives on various collection media, coated with DNPH with 1,3-butanediol, stored at room temperature for (a) 1 and (b) 7 d (N ) 6 per pad type). crotonaldehyde. In a subsequent experiment with DNPH/1,3-butanediol solution, glass fiber and Teflon-coated glass fiber pads were retested along with two additional media, C18 silane-treated silica-celluose pads (C18-SS), and C18 silica TLC plates (C18TLC). After spiking with aldehyde-DNPH derivatives and storing for 24 h at room temperature, recoveries were close to 100% for formaldehyde-, acetaldehyde-, glyoxal-, and methylglyoxal-DNPH derivatives on most media (Figure 6a). Glass fiber pads showed the best recoveries, especially for acrolein and crotonaldehyde. On day 7, glass fiber pads again showed the best recoveries (near or above 100%) for most aldehydes except for acrolein (68%) and glyoxal (132%) (Figure 6b). Note that glyoxal concentration increased from day 1 to day 7 on all media except for C18-TLC. While the recovery of glyoxal and methylglyoxal remained near or above 100% on all media on day 7, recovery of other aldehydes on day 7 on Teflon-coated glass fiber, C18 silane-treated silicacellulose pads, and C18-silica TLC plates was lower than that on the glass fiber pads. To test our hypothesis about the catalyzed reactions due to the acidity in the original DNPH derivatization solution, we substituted citric acid/sodium citrate buffer (pH 4) for the hydrochloric acid used in the original solution. In light of the above observations, 1,3-butandiol was substituted for glycerol, and 12 replicate glass fiber pads were used as the collection medium. The recovery of aldehyde-DNPH derivatives, especially acrolein and crotonaldehyde derivatives, improved from those shown in Figure 6, ranging between 88% and 100% on day 1 and between 89% and 105% on day 7, respectively (Table 2). Thus, improved recovery (above 90%) for acrolein and crotonaldehyde was attained for up to 7-d storage on glass fiber pads. Optimization of Coating Agents. The above results validated our hypotheses about the sampling medium, the hygroscopic agent, and the acid-catalyzed reaction of acrolein- and crotonaldehyde-DNPH derivatives. Optimization tests were subsequently conducted to determine the 2306

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acrolein

crotonaldehyde

glyoxal

methylglyoxal

buffer

pH

buffer concn (mM)

H/P C/S C/S C/S H/P C/S C/S C/S H/P C/S C/S C/S H/P C/S H/P C/S C/S H/P C/S H/P C/S H/P C/S C/S H/P H/P C/S C/S C/S H/P

1 4 4 3 1 4 4 3 1 4 4 4 1 3 1 4 4 1 3 1 4 1 4 3 1 1 4 3 3 1

10 10 20 10 10 20 10 10 10 10 20 10 10 10 10 20 10 10 10 10 10 10 10 10 10 10 10 10 10 10

diol concn (%)

collection efficiency (%) (N ) 3)

0.4 10 1 10 1 1 0.4 10 1 1 1 0.4 1 10 0.4 1 0.4 0.4 10 1 10 0.4 5 0.4 1 0.4 10 0.4 10 1

57 ( 25 55 ( 26 49 ( 16 44 ( 6 43 ( 17 87 ( 7 56 ( 12 54 ( 3 44 ( 28 42 ( 4 56 ( 8 29 ( 10 23 ( 20 21 ( 1 18 ( 4 61 ( 9 32 ( 10 26 ( 6 22 ( 1 22 ( 14 46 ( 41 38 ( 27 31 ( 23 26 ( 5 26 ( 27 31 ( 18 30 ( 21 25 ( 9 24 ( 7 20 ( 15

a Calculated collection efficiency was the aldehyde concentration measured in the pad relative to the aldehyde concentration in the spiking solution. H/P represents hydrochloric acid with potassium chloride; C/S represents citric acid with sodium citrate.

optimal pH value and concentrations of the buffer solution and 1,3-butanediol in the DNPH derivatization solution. These tests were performed by exposing coated glass fiber pads to un-derivatized aldehydes inside a sealed Tedlar bag. Results (Table 3) from the first optimization experiment indicated that citric acid/sodium citrate (C/S) buffer (pH 4) provided the highest collection efficiency for most aldehydes, while hydrochloric acid/potassium chloride (H/P) buffer (pH 1) provided the best collection efficiency for formaldehyde and methylglyoxal. Overall, the highest recoveries were given by the C/S buffer at pH 4. The derivatization solution with the 20 mM C/S buffer provided one of the top three best collection efficiencies for four aldehydes. The collection efficiency for most aldehydes was the lowest at C/S concentration of 100 mM at pH 4, as compared to 10 and 20 mM of C/S at pH 4. This indicated that the optimal buffer concentration was between 10 and 100 mM. Phosphoric acid

FIGURE 7. Collection efficiencies of aldehyde-DNPH derivatives on glass fiber pads coated with various butanediol and citric acid/sodium citrate buffers (N ) 5 per symbol).

FIGURE 8. Collection of aldehyde-DNPH derivatives on glass fiber pads coated with 50 mM citric acid/sodium citrate buffer (pH 4) and three levels of butanediol concentrations (N ) 6 per box plot). solutions (results not shown) destroyed most unsaturated aldehydes and showed the lowest recovery for formaldehyde and acetaldehyde on glass fiber pads. The second optimization test was to optimize concentrations of the C/S buffer at pH 4 and 1,3-butanediol. Recoveries increased with buffer concentration for most aldehydes, except for glyoxal and methylglyoxal (Figure 7). For most aldehydes, the 50 mM buffer provided the best collection efficiency. The collection efficiency also increased with butanediol concentration between 0.1 and 10%, peaked at 10%, and dropped abruptly at 20% for most aldehydes. The

exceptions were glyoxal and methylglyoxal. The collection efficiency for glyoxal and methylglyoxal was affected by an interaction between the buffer and butanediol concentrations. At 50 mM buffer solution, the peak collection efficiency was observed at 5% of butanediol for glyoxal, while the highest collection efficiency for methylglyoxal for both 20 and 50 mM buffers was observed at 5% butanediol. In summary, the optimal combination was either 5% or 10% of butanediol with 50 mM citric acid/sodium citrate buffer at pH 4. The last optimization experiment tested derivatization solutions at 50 mM buffer and 5, 10, and 20% of butanediol VOL. 35, NO. 11, 2001 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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concentrations. The highest collection was observed at 10% butanediol for formaldehyde, acetaldehyde, acrolein, and crotonaldehyde (Figure 8). For glyoxal, collection efficiency was not significantly different between the solutions with 10 and 20% butanediol. For methylglyoxal, recovery efficiency at 20% butanediol was significantly higher than that at either 5% or 10% butanediol. Thus, a new formulation of the DNPH derivatization solution with 50 mM C/S buffer at pH 4 and 10% butanediol should provide the optimum collection efficiency for the six target aldehydes over a 7-d sampling period.

Acknowledgments This study was supported by the Health Effects Institute (Project No. 98-3) and, in part, by the Department of Environmental Health, University of Washington. We thank Ms. Hongbin Xiao for her assistance in this research project. We thank Dr. Kochy Fung of AtmAA, Inc. and Dr. Debra Kaden of HEI for their advice. We are grateful to Dr. Charles W. Gardner, chief scientist at Bacharach, Inc., for his technical support, advice, and GDM filter supplies.

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Received for review October 20, 2000. Revised manuscript received February 22, 2001. Accepted March 5, 2001. ES001795C