Envlron. Sci. Technol. 1986, 20, 11 17-1 122
Chlorination Byproducts of Amino Acids in Natural Waters Michael L. Trehy,"It Rlchard A. Yost,+ and Carl J. Miles$ Chemistry Department and Pesticide Research Laboratory, University of Florida, Gainesville, Florida 326 11
rn The chlorination of aspartic acid, tyrosine, and tryptophan was studied at acidic and neutral pH by gas chromatography/mass spectrometry. Tandem mass spectrometric analysis of chlorinated tyrosine provided additional structural information necessary for the identification of the chlorination intermediates. The amino acids were found to form dichloroacetonitrile and chloral as major byproducts. The formation of chloral was found to significantly increase as the pH was raised from 7 to 8, and the pH dependence is consistent with the reaction pathways proposed. Chlorinated water samples from a natural lake and from a wastewater treatment plant were found to contain significant amounts of both chloral and dichloroacetonitrile. Chlorination of natural waters has been shown to result in the formation of halogenated byproducts (1,2). Humic and fulvic substances present in natural waters react rapidly with free chlorine to form trihalomethanes (THM) and other chlorinated and nonchlorinated byproducts (2-8). Free, proteinaceous and humic-bound amino acids are also potential precursors for chlorination byproducts (9-13). Amino acids are ubiquitous constituents of the environment (14-18). Analyses of water and sediment samples for total amino acid content following hydrolysis of any proteinaceous matter have shown that amino acids are present at significant concentrations in such diverse water sources as the following: the coastal plain rivers of Southeastern United States, 1.35-2.74 ppm with aspartic acid having an average concentration of 270 ppb (15);the Williamson River (Oregon), approximately 1 ppm with aspartic acid having an average concentration of approximately 150 ppb (16); waters of the Mackenzie River drainage basin (Canada), 12-299 ppb (14); domestic wastewaters, 0.34-2.0 ppm with aspartic acid varying from 20 to 130 ppb (18). The relative concentration of the condensed amino acids to free amino acids has been found to vary from 2 to 15 in water samples from the Mackenzie water drainage district (14). The chlorination of amino acids and peptides has been studied (14;28). These studies indicate that amino acids react with an equimolar amount of halogenating agent to yield the corresponding aldehyde. However, if an excess of halogenating agent is added, then the corresponding nitrile can also be formed, with the ratio of the aldehyde to nitrile formed increasing with pH (24). In the case of alanine, for example, the amino acid is oxidized to acetonitrile and acetaldehyde. Subsequently or concomitantly to oxidation the amino acids can also be chlorinated. Dichloroacetonitrile has been shown to be formed on chlorination of aspartic acid, tyrosine, and tryptophan (9, 10);however, the formation of chloral was only speculated. Some aromatic amino acids such as tyrosine undergo haloform reactions similar to the reaction pathways determined for humic sustances. Substitution of chlorine on the aromatic ring of tyrosine has been demonstrated at pH 1-2 (25). *Address correspondence to this author at the Monsanto Co., 800 N. Lindbergh, St. Louis, MO 63167. Chemistry Department. t Pesticide Research Laboratory. 0013-936X/86/0920-1117$01.50/0
Dihaloacetonitriles (6, 10-12, 29) and chloral hydrate (30) have been found in chlorinated natural waters. Although chloral can be formed as a result of chlorination of humic substances (3, 5, 7, 81, the presence of dihaloacetonitriles (DHANs) in chlorinated waters strongly implicates amino acids or other proteinaceous material as playing a significant role in the formation of chlorination byproducts. The chlorination byproducts of amino acids are of concern since dichloroacetonitrile has been found to be mutagenic by the Ames test (31,32)and tumor-initiating by the Sencar mouse skin bioassay (33). In addition it has been shown that consumption of chlorinated water results in the in vivo formation of dichloroacetonitrile and trichloroacetonitrile (13,34). The potential for in vivo formation of chlorination byproducts should be considered, since chlorine has been shown to react with polypeptides (22) as well as with amino acids. Monochloroamines, which are used as an alternative disinfectant for free chlorine, decompose to form free chlorine at low pH (35). The use of combined chlorine for the reduction of chlorination byproducts may not be an effective means for lowering the formation of in vivo chlorination byproducts. In this paper the formation of chlorination byproducts of the amino acids aspartic acid, tyrosine, and tryptophan is investigated. Experimental Section Purge and Trap (PT). A 10-mL aliquot of sample or standard solution was transferred to a 25-mL fritted VOA sampler tube (SSIScientific Products; Micanopy FL)and attached to a PT device constructed by using EPA protocol (36). This solution was purged with helium at about 65 mL/min for'7 rnin onto a 0.2 cm X 25 cm Tenax trap (80/100 mesh). Approximately 1 min before the purge was complete, a small loop of the capillary GC column (0.2 mm i.d. X 30 M DB5,0.25 pm film thickness; J & W Scientific, Cordova, CA) near the injection port was immersed into a cup of liquid nitrogen. After the 7-min purge, the trap was heated to 180 "C, and the Tenax trap effluent was directed onto the capillary column and into the liquid nitrogen focus trap using a 1:lO split injection. After 4 min of heating at 180 OC, the Tenax effluent was switched off, capillary column flow restored to 2 mL/min He, and the liquid nitrogen removed. After an initial hold of 1 min at 30 "C, the column was temperature programmed to 280 "C at 10 OC/min. Following the start of the GC run, the mass spectrometer (Hewlett-Packard 5985B) was continuously scanned from 45 to 200 amu at 800 amu/s (5 scans/s). Positive electron impact spectra were collected at a source temperature of 200 "C and an electron multiplier voltage of 2000 V. After each run the column was baked out at 280 "C for at least 10 min to remove the late-eluting peaks. The Tenax trap was also baked out at 180 " C for an additional 5 min to remove low voltaility compounds. All glassware used in this method was baked in an oven at 200 "C for at least 30 rnin prior to use. Liquid-Liquid Extraction (LLE). Aqueous samples were collected in 60-mL septum vials which contained a magnetic stirring bar and were extracted with rapid stirring for 15 min with 15 mL of either pentane or ethyl ether. Pentane did not extract chloral hydrate from aqueous samples. However, ethyl ether was found to be suitable for the determination of chloroform, dihalocetonitriles,and
0 1988 American Chemical Soclety
Environ. Scl. Technol., Vol. 20, No. 11, 1986
1117
chloral hydrate. Ethyl ether extracts were dried by passing the extract through a Pastuer pipet packed with anhydrous sodium sulfate. Determination of chloroform, dichloroacetonitrile, and chloral hydrate was carried out by injecting 3 CLLof the dried extract onto a 6 f t X 2 mm i.d. glass column packed with 5% diethylene glycol succinate on Chromosorb W/AW 80/100 mesh (Supelco) with helium as carrier gas. The injection port was operated at 180 O C , and the temperature program which separated chloral hydrate from dichloroacetonitrile was a 40 OC initial temperature for 1min, 25 OC/min to 65 "Cand hold for 5 min, and finally 15 OC/min to 150 "C. A 25-m DB5 0.2 mm i.d. open tubular column (J & M) did not resolve dichloroacetonitrile from chloral hydrate, although this column offered a considerable improvement in sensitivity and peak shape. A Carbowax 20M packed column (Supelco) gave very poor limits of detection for chloral hydrate. Quantitative analysis of the ethyl ether extracts was carried out in the negative chemical ionization (NCI) mode. Selected ion monitoring for the chloride 35 and 37 ions was employed for maximum sensitivity with the electron multiplier set at 950 V. Chlorination of Amino Acids. Stock chlorine solutions were prepared by bubbling gaseous chlorine (Matheson) through 200 mL of deionized water containing approximately 100 mg of NaOH. Chlorine stock solutions were determined to be approximately 2000 ppm by the DPD titrimetric procedure (37). The reaction of aspartic acid, tyrosine, and tryptophan was studied at ambient temperature at pH 1-2 in order to investigate the formation of chlorination intermediates (27)and at neutral pHs to determine byproducts likely to be formed on chlorination of natural waters. No effort was made to exclude room light from the samples. Samples were dechlorinated with excess sodium sulfite (approximately 1O:l mole ratio of sodium sulfite to chlorine added) in order to avoid contamination with elemental sulfur formed on dechlorination with sodium thiosulfate. Analysis of samples for brominated analogues would be improved by dechlorinating with thiosulfate since sulfite has been found to react with the brominated DHANs (9). Apparatus. Analysis of samples by purge and trap was carried out with Hewlett-Packard 5985B mass spectrometer while analysis for samples analyzed by liquid-liquid extraction was carried out on either a Finnigan MAT TSQ45 or on a Finnigan MAT 4515. A Finnigan MAT TSQ45 gas chromatograph/triple-stage quadrupole mass spectrometer/data system was used for the determination of the daughter ion mass spectra and for the quantitative analysis of the ethyl ether extracts of environmental samples and chlorinated amino acids at low concentrations. A Finnigan MAT 4515 GC/MS was employed for the analysis of the ethyl ether extracts of chlorinated amino acids at high concentrations. Electron energies of 70 and 100 eV were used for electron ionization (EI) and chemical ionization (CI), respectively. The chemical ionization reagent gas was methane or isobutane at an ionizer pressure of 0.8 Torr. The mass spectrometer was tuned with FC43 (perfluorotributylamine). Nitrogen collision gas was introduced at 1.7 mTorr at collision energies of 10-20 eV for collision-induced dissociation (CID).The continuous dynode electron multiplier was operated at 850-950 V, with the conversion dynodes at &3000V. The preamplifier gain was set at los VIA. The open tubular columns were inserted directly into the ion source. When packed columns were used, a glass jet separator was employed. Discussion of Analytical Techniques
Determination of chloroform, dichloroacetonitrile, and 1118 Environ. Scl. Technol., Vol. 20, No. 11, 1986
Table I. Extraction Efficiency and Limit of Detection (LOD) in ppb Determined for Chloroform, Dichloroacetonitrile, and Chloral Hydrate by Purge and Trap by GC/MS in E1 Mode with Full Scan vs. LLE with Diethyl Ether by GC/MS in NCI Mode with SIM for the Chlorine m /z35 Ion ethyl ether extraction LOD efficiency CHCls CHClpCN CC13CH(OH)2
purge and trap efficiency
LOD
91 100 36
1 1 4
1 17
91 16
c5
Table 11. Response for Chloroform and Dichloroacetonitrile vs. Purge Time' time, min
chloroform
3.5 7.0 14.0 28.0
13 326 22 597 10 371 4041
areas dichloroacetonitrile 622 1429 1777 7085
chloral hydrate was evaluated for the PT technique and for LLE with pentane and with ethyl ether. Preliminary investigations demonstrated that chloral hydrate was not extracted from water with pentane. However, chloral hydrate can be extracted from water with ethyl ether and has been found to have a partition coefficient of approximately 4 (38). The extraction efficiency for chloral hydrate obtained when the ratio by volume of ethyl ether to water was one to five after mixing was 36%, which corresponds to a partition coefficient of 3 at a concentration of 200 ppb in deionized water (Table I). Dichloroacetonitrile and chloroform were extracted from deionized water with extraction efficiences of 100% and 91% ,respectively, with ethyl ether. Comparison of the PT and the LLE methods using standard solutions showed similar limits of detection (LOD) for chloroform, while for dichloroacetonitrile the LOD was 17 times better for the LLE technique than for the PT technique and chloral hydrate could not be detected by the PT technique (Table I). Although the difference in the LOD is partidy attributable to the different ionization methods used, the major difference in the LOD observed is due to the low purge efficiency for dichloroacetonitrile and chloral hydrate. To evaluate the purge efficiency, a sample of deionized water was spiked with 119 ppb of chloroform, 110 ppb of dichloroacetonitrile, and 85 ppb of chloral hydrate and was purged with helium a t a rate of 70 mL/min. The response for dichloroacetonitrile increases linearly with increasing purge time during the initial 28 min, indicating that a substantial portion of the original dichloroacetonitrile still remains in solution even after 28 min (Table 11). The loss of chloroform from the trapping column at a purge time greater than 7 min demonstrates the inability of the purge and trap technique to simultaneously determine compounds with diverse volatilities and aqueous solubilities. The brominated members of the dihaloacetonitrilesare less volatile and more difficult to determine by the PT method than dichloroacetonitrile (12). These resulta indicate that the PT technique will not detect low concentrations of dichloroacetonitrile or chloral hydrate in water samples and &at an alternative technique such as LLE is needed. The determination of the dihaloacetonitriles (DHANs) is further complicated by the reaction of the DHANs with common dechlorinating agents and hydrolysis in the pH
400,
P t 7
HOC CH2$ COH NX
P
4
HOCCHzCHO
LFI
1 /,
c I 3~ H (OHl2
/ C'
HOCCH2CN
0 CX,CHO
CHX2CN
0
Flgure 1. Chlorination of aspartic acid.
range from 7 to 9 (9). At pH 8.3 the half-lives (in hours) for the DHANs in the presence of sodium thiosulfate and in its absence are the following: dichloroacetonitrile, 25 and 29; bromochloroacetonitrile, 32 and 55; dibromoacetonitrile, 12 and 85. The rate of degradation of the DHANs by sodium sulfite is even greater, so that the brominated members cannot be detected 40 min after dechlorination, while only a 10% loss of dichloroacetonitrile was observed after 24 h (9). The brominated members of the trihaloacetaldehyde (chloral) series were not investigated. The determination of chloral hydrate is difficult due to the conversion of chloral hydrate to chloral on heating. When chloral hydrate is extracted from water with ethyl ether and injected onto the column, the relatively nonvolatile and more polar chloral hydrate is strongly retained by the column. However, as the column temperature is programmed to higher temperatures, conversion of chloral hydrate to chloral occurs and results in peak broadening. Analysis with the 25 m X 0.25 mm i.d. DB5 thick-film open tubular column (SE54 equivalent) gave the best results for peak shape, but dichloroacetonitrile and chloral hydrate eluted at essentially the same time. Thus, when selected ion monitoring for the m / z 35 ion in NCI by GC/MS is employed, it is difficult to quantitate these two components separately. A 25 m X 0.25 mm i.d. SP20 open tubular column (Carbowax 20 M equivalent) was also evaluated. The SP20 column resulted in substantial peak broadening for chloral hydrate. Several packed columns were also evaluated. Of these, a 10% diethylene glycol succinate (DEGS) column was found to give the best results while squalane and Carbowax 20 M packed columns gave poor results. The capability of the mass spectrometer to discriminate between different ions greatly aided in the selective detection of chlorination byproducts. Reactions with Aqueous Chlorine Aspartic Acid. Aspartic acid is second only to glycine in occurrence in natural waters (14)and is known to form dichloroacetonitrile when chlorinated (9,lO). Aspartic acid is also of interest since the reaction pathway suggested for aspartic acid indicates that not only dichloroacetonitrile but also chloral should be formed, as shown in Figure 1. In order to investigate this reaction, a 10 ppm solution of aspartic acid was chlorinated at pH 6.4 with 33 ppm of chlorine (chlorine to amino nitrogen ratio was approximately 6). The free chlorine residual after 162 min was 7.6 ppm, indicating that an excess of chlorine was present. Aliquots of the chlorinated water were dechlorinated with sodium sulfite immediately prior to extraction with ethyl ether. As had been anticipated, aspartic acid was found to yield not only dichloroacetonitrile but chloral as well. After 162
15 TIME
30
iminl
Flgure 2. Rate of conversion of aspartic acid to chloral and dichloroacetonltrlle.
min the 10 ppm solution of aspartic acid yielded 0.57 ppm (6.9% yield) of dichloroacetonitrile and 0.33 ppm (2.7% yield) of chloral hydrate. The rate of conversion of aspartic acid to chloral and dichloroacetonitrile on chlorination at pH 6.4 for the first 30 min is shown in Figure 2. Increasing the pH at which the chlorination of amino acids occurs has been shown to favor the formation of the aldehyde (24). In order to investigate the influence of pH on chlorination of aspartic acid, an 80 ppm solution of aspartic acid was chlorinated at pH 8.5 with an initial chlorine dose of 280 ppm (chlorine to amino nitrogen ratio of approximately 7) and dechlorinated with sodium sulfite immediately prior to extraction with ethyl ether. After 1h the aspartic acid solution was found to contain 0.4 ppm (0.6% yield) of dichloroacetonitrile and 3.5 ppm (3.5% yield) of chloral hydrate. Although a comparison of the relative rates for the reaction of chlorine with amino acids at different pHs requires further investigation, it appears that the relative ratio of the byproducts is greatly influenced by the pH at which the reaction occurs, with a much greater abundance of chloral forming at the elevated pH. Chloroform was not formed during the chlorination step but appears to result entirely as a result of the relatively slow hydrolysis of chloral. The half-life of chloral at pH 8 and 35 OC is 2 days (38). Tyrosine and Tryptophan. The chlorination of tyrosine at pH 1-2 (26) has been reported to result in the formation of mono- and dichloro 4-hydroxybenzeneacetonitrile (I) and mono- and dichloro-4-hydroxybenzeneacetaldehyde (11). The E1 mass spectra of the H
CHZC N e
o
(I)
cH,cHo
H0-Q(
n,
chlorination byproducts of tyrosine have been interpreted (26)to be consistent with chlorination of the aromatic ring rather than of either the aldehyde or aliphatic nitrile portion of the molecule. In order to investigate this reaction a 74 ppm solution of tyrosine was chlorinated at pH 2 with 1200 ppm of chlorine and dechlorinated with sodium sulfite after l h. The sample was extracted with ethyl ether, dried with anhydrous sodium sulfate, and analyzed by GC/MS employing electron ionization (EI), positive chemical ionization (PCI), and negative electroncapture chemical ionization (NCI) with methane as the reagent gas. The reconstructed ion chromatogram for the PCI analysis of the ethyl ether extract by GC/MS employing a 5-m DB5 thick-film open tubular column showed three major peaks with chlorine isotope patterns (Figure 3). The Environ. Sci. Technol., Vol. 20, No. 11, 1986
1119
5
GC/MS
2' H0
N
202
-
175
i rnlr
4
6
6
TI ME
162
50
I50
100
200
250
Flgure 5. Daughter spectrum obtained for peak 2' during the GC/ MSIMS analysis.
(mlnl
Flgure 3. Reconstructed ion chromatogram for the PCI analysis of the ethyl ether extract of chlorinated tyrosine. I66
2'
I
201 CX3CH0
166
I
or
2
I
CHX,
I
m/z
r 100
I40
180
220
Flgure 4. Chlorine isotope patterns.
chlorine isotope patterns shown in Figure 4 indicate that peak 1 is due to a compound that contains at least one chlorine while peaks 2 and 2' appear to contain at least two chlorines. The EI, PCI, and NCI mass spectra are consistent with the formation of mono- and dichloro-4hydroxybenzeneacetonitrile. Although the mass spectra are consistent with chlorination of the aromatic or aliphatic portion of the molecule, the location of the chlorines could not be conclusively determined. In order to identify the location of the chlorines, tandem mass spectrometry was used in the daughter ion mode. The PCI ion m / z 202, peak 2', was mass-selected in the first quadrupole and fragmented in the center quadrupole by collisions with nitrogen, and the fragment ions formed were then mass-analyzed in the third quadrupole. The daughter spectrum obtained for peak 2' during the GC/ MS/MS analysis (shown in Figure 5) indicates that the chlorines are located on the aromatic ring, as has been proposed by Burleson et al. (26). The loss of 40 amu from the m l z 202 ion to give the m/z 162 ion can be explained by the loss of CH2CN. This fragmentation is consistent with chlorination on the aromatic ring. The formation of chlorinated 4-hydroxybenzeneacetaldehydewas not detected. Volatile chlorination byproducts of tyrosine and tryptophan were determined by ethyl ether LLE and by the PT technique. Tyrosine and tryptophan were found to form chloroform, dichloroacetonitrile,and chloral hydrate. Chloral hydrate was only detected by the ethyl ether extraction procedure. At pH 8.5 a 45.6 ppm solution of tyrosine was chlorinated for 1h with 280 ppm of chlorine 1120 Environ. Sci. Technol., Vol. 20,
No. 11,
1986
Flgure 6. Chlorination of tyroslne.
(chlorine to amino acid ratio was approximately 16) and yielded 0.31 ppm of chloroform, 0.019 ppm of chloroacetonitrile, and 0.23 ppm of chloral hydrate. A 58.4 ppm solution of tryptophan (chlorine to amino acid ratio was approximately 14) yielded 5.2 ppm of chloroform,0.23 ppm of dichloroacetonitrile, and 6.2 ppm of chloral hydrate in 1 h. The reaction pathways for tyrosine with aqueous chlorine, shown in Figure 6, suggest that unlike aspartic acid, both tyrosine and tryptophan form chloroform directly. The formation of chloroform quite likely occurs by chlorination of the aromatic ring with subsequent ring opening and hydrolysis, while the formation of chloral and dichloroacetonitrile occurs by chlorination of the amino group and subsequent oxidation and further chlorination. Formation of Chloral Hydrate and Dichloroacetonitrile in Environmental Samples
The presence of free and condensed amino acids in natural waters (14-18) and known reaction pathways for the chlorination of amino acids (19-28) strongly indicate that amino acids are potential precursors in the environment for chlorination byproducts. The rate of formation of chlorination byproducts is expected to be greatly influenced by the form the amino acids are present in since the amide group of the peptides will react much more slowly than will the amino group of the free amino acid (39). In order to investigate the formation of dichloroacetonitrile and chloral hydrate in environmental samples, two different lakewater samples were chlorinated, and the chlorinated effluents from two domestic wastewater treatment plants were collected and analyzed for chlorination byproducts. Samples of chlorinated wastewater from an extended aeration treatment plant collected on two separate days were analyzed and found to contain 32-80 ppb of chloroform, 7-14 ppb of dichloroacetonitrile, and 20-38 ppb of chloral hydrate. The reconstructed ion chromatogram for one of the samples from the extended aeration plant is
0
I
Table 111. Concentration of Chloroform, Dichloroacetonitrile, and Chloral Hydrate Formed with Time of Contact with Chlorine for Lake Water Samples Collected in Gainesville, FL ( G ) ,and West Palm Beach, FL (W), at pH 7.2 and 7.6, Respectively
I( CHCI,CN
I
I\ T I ME
contact chloroform, dichloroacetonitrile, chloral, sample time, min ppb PPb PPb 61 10 9 G 10 88 13 23 G 30 108 17 35 G 50 116 17 35 G 70 W 5 126 5 8 163 14 12 W 15 239 19 30 W 70
(mln)
Flgure 7. Reconstructed ion chromatogram for one of the samples from the extended aeration plant.
shown in Figure 7. Purge and trap analysis of the wastewater sample confirmed the concentration and identity of chloroform and dichloroacetonitrile in the chlorinted wastewater sample. The E1 mass spectrum and retention time matched those of dichloroacetonitrile. The presence of chloral was confirmed by extracting 1 L of chlorinated wastewater effluent first with 200 mL of ethyl ether and then a second time with 100 mL, combining the extracts and drying with anhydrous sodium sulfate, and concentrating to 1 mL with a stream of nitrogen. This sample was then analyzed by positive chemical ionization selected ion monitoring mode for the major ions of chloral hydrate, m/z 71,73,110, and 112. The retention time and ratio of these ions matched those for chloral hyrate. Samples from the wastewater treatment plant employing split treatment with activated sludge and trickling filters did not contain detectable concentrations of chlorination byproducts. The presence of ammonia in the effluent of the trickling filters is quite likely responsible for the absence of chlorination byproducts, since chlorine rapidly combines with ammonia to form chloramines which form chlorination byproducts slowly if at all (40). Analysis of chlorinated lakewater ‘samples also demonstrated the formaton of both dichloroacetonitrile and chloral hydrate. A sample of lake water from the West Palm Beach, FL, area was chlorinated at pH 7.2 for 70 min with an initial dose of 12 ppm of chlorine and produced 239 ppb of chloroform, 19 ppb of dichloroacetonitrile, and 30 ppb of chloral hydrate. Lake water collected in the Gainesville, FL, area was chlorinated at pH 7.6 with an initial dose of 20 ppm of chlorine and was found to contain 116 ppb of chloroform, 17 ppb of dichloroacetonitrile, and 35 ppb of chloral hydrate after 70 min. The rate of formation of chloroform, dichloroacetonitrile, and chloral hydrate on chlorination of the lake water samples is shown in Table 111. Although both lake waters had rather similar formation potentials for dichloroacetonitrile and chloral hydrate, the formation potential for chloroform appears to be quite different. The independence of the chloroform formation potential with respect to the dichloroacetonitrile and chloral hydrate formation potentials is indicative of different precursors for these chlorination byproducts, humic substances vs. amino acids or other nitrogenous substances. Conclusions
In this study the formation of chloroform, dichloroacetonitrile, and chloral hydrate upon chlorination of the amino acids aspartic acid, tyrosine, and tryptophan has been demonstrated. The purge and trap method did not recover chloral hyrdate and recovered dichloroacetonitrile
inefficiently, indicating that this technique is inappropriate for the determination of these compounds in water. The presence of dichloroacetonitrile and chloral in chlorinated natural waters strongly implicates amino acids and other nitrogenous compounds as potential precursors in natural waters. Like humic substances, amino acids abd other proteinaceous substances are ubiquitous to the environment and should be considered in the development of environmental policies regarding chlorination byproducts. Acknowledgments
Special thanks are due to Theodore 1. Bieber for his valuable discussions and to David Berberich for technical assistance. Registry No. Aspartic acid, 56-84-8;tyrosine, 60-18-4; tryptophan, 73-22-3; dichloroacetonitrile, 3018-12-0; chloral, 75-87-6; chloroform, 67-66-3.
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fication and Analysis of Organic pollutants in Water; Keith, L. H., Ed.; Ann Arbor Science: Ann Arbor, MI, 1976; pp 329-373. (31) Simmon,V. F.; Kauhanen, K.; Tardiff, R. G. Dev. Toxicol. Environ. Sci. 1977, 2, 249-258. (32) Bull, R. J. In Water ChlorinationEnvironmental Impact and Health Effects;Jolley, R. L.; Brungs, W. A.; Cotruvo,
J. A.; Cumming, R. B.; Mattice, J. S.; Jacobs, V. A., Eds.; Ann Arbor $cience: Ann Arbor, MI, 1983; pp 1401-1415. (33) Bull, R. J.; Robinson, M. In Water Chlorination: Chemistry, Environmental Impact and Health Effects; Jolley, R. L.; Bull, R. J.; Davis, W. P.; Katz, S.; Roberts, M. H., Jr.; Jacobs, V. A., Eds.; Lewis Publishers: Chelsea, MI, 1985; V O ~5,. pp 221-228. (34) Mink, F. L.; Coleman, W. E.; Munch, J. W.; Kaylor, W. H.; Ringland, H. P. Bull. Environ. Contam. Toxicol. 1983,30, 394-399. (35) Palin, A. T. Water Water Eng. 1950, 53, 189-200. (36) Fed. Regist. Part VIII, Method 624, Friday Oct 26, 1984, Washington, DC, pp 141-152. (37) Standard Methods for the Examination of Water and Wastewater,16th ed.; American Public Health Association: Washington, DC, 1985. (38) Luknitskii, F. 1. Chem. Rev. 1975, 75, 259-289. (39) Morris, J. C. In Principles and Applications of Water
Chemistry;Haust, S. D.; Hunter, J. W., Eds.; Wiley: New York, 1967; pp 23-53. (40) Stevens, A. J. Am. Water Works Assoc. 1976,68,615-620. Received for review November 15, 1985. Revised manuscript received April 14,1986. Accepted May 12,1986. This work was supported by a grant from the U.S. Environmental Protection Agency under Contact 810973-01-0.
Regional Tree Growth Reductions due to Ambient Ozone: Evidence from Field Experiments Deane Wang” and F. Herbert Bormann Yale School of Forestry and Environmental Studies, New Haven, Connecticut 0651 1
David F. Karnosky School of Forestry, Michigan Technological Universlty, Houghton, Michigan 4993 1
rn Observations from extensive regions in Europe and North America suggest that many forests may be in early stages of ecosystem decline. We present experimental evidence from open-top chamber field studies indicating that ambient ozone at levels below the ambient air quality standard (235pg m-3) causes significant reductions (19%) in the growth of sapling poplars (hybrid Populus). While ozone-induced reductions in growth have been observed under laboratory and greenhouse conditions, demonstration of this effect under field conditions is critical to the establishment of ozone standards. Growth reductions for Populus deltoides and Robinia pseudoacacia were not significant (CY = 0.05). Reductions in productivity and height growth occurred without visible symptoms of foliar injury and at ozone concentrations below current standards. If this “invisible” injury is typical in other tree species, the extent of ozone-induced forest damage may presently be greatly underestimated. Additional field studies on a regional basis are needed. Introduction
As attention on the causes of widespread forest decline *Address correspondence to this author at the Center for Urban Horticulture GF-15, University of Washington, Seattle, WA 98195. 1122 Environ. Scl. Technol., Vol. 20, No. 11, 1986
(1-5) shifts from SOz to NO, and its photochemical products such as ozone (6),it becomes increasingly important that scientific observations documenting ozone dose response be determined prior to undertaking major policy initiatives to deal with the problem. Field experiments have documented regional growth reductions in agricultural crops (IO),but forest tree responses to ambient ozone have not been adequately studied. Recent laboratory research (7) has provided one critical link connecting low, ambient levels of ozone and reduction of net photosynthesis in trees. Past work has shown important inconsistencies in pollutant dose response of plants under laboratory and greenhouse conditions as compared to response under field conditions (8, 9). Thus, field experiments are essential, providing realistic calibration points for dose-response functions which can be elucidated in more detail in the laboratory. Field research is limited by experimental artifacts including chamber effects and artificial plant assemblages. In addition, field trials have generally involved fumigations (10). These ozone additions have often been introduced abruptly, causing an initial rapid change in ozone concentrations. Also, fumigated chambers are subject to rapid wind-induced fluctuations in chamber concentrations (Figure 1). The effects of these conditions on dose re-
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