Role of Amide Nitrogen in Water Chlorination: Proton NMR Evidence

In the first case, Cl would displace the amide H, and electron withdrawal by Cl+ .... Conditions: p11 echo water suppression, 256 scans, 298 K. Peak a...
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Environ. Sci. Technol. 1999, 33, 3568-3573

Role of Amide Nitrogen in Water Chlorination: Proton NMR Evidence JAMES S. JENSEN,* YIU-FAI LAM, AND GEORGE R. HELZ Department of Chemistry and Biochemistry and Water Resources Research Center, University of Maryland, College Park, Maryland 20742

A large fraction of the dissolved amino-N in natural waters and wastewaters is contained in amide groups, for example, in proteins. Whether this N pool reacts with and modifies the chemical behavior of chlorine during water disinfection is unclear. To investigate this issue, watersuppressed, proton NMR spectra have been obtained for aqueous, acetylated glycine, alanine, and alanylalanylalanine before and after treatment with NaOCl at near-neutral pH. N-Chlorination of N-acetylglycine (aceturic acid) induced cis-trans rotation about the amide bond at ambient temperature. N-Bound Cl+ induced a measurable downfield chemical shift in the acetyl methyl resonance in N-acetylglycine, ∂δ ) 0.15 ppm (cis) and ∂δ ) 0.3 ppm (trans), and in N-acetylalanine methyl resonances, acetyl methyl ∂δ ) 0.3 ppm (trans) and side chain methyl ∂δ ) 0.2 ppm. Chlorination of N-acetylalanylalanylalanine produced ∂δ values similar to N-acetylalanine. The spectral effects were reversible, the original spectra being regenerated upon dechlorination with sulfite. Negligible substrate decomposition was observed. Rate constants for chlorination of N-acetylalanine near neutral pH are kf ) 1.58 × 10-3 M-1 s-1 and kb ) 7.57 × 10-7 s-1 where Keq ) 2.1 × 103. Because of both sluggish formation kinetics and an unfavorable equilibrium constant, N-chloramides are predicted to be unimportant under typical disinfection conditions.

Introduction Disinfection of drinking water and wastewater with Cl2 cannot be understood without understanding the role of amino nitrogen. Both Cl2 (aq) and its hydrolysis product, HOCl, react quickly with organic and inorganic amino-N, producing chloramines (1, 2). The chloramines occupy a key position in the disinfection process because they are temporary reservoirs of disinfection capacity, both moderating and sometimes prolonging disinfection capability (3). They are also intermediates in reactions leading to undesirable byproducts such as cyanogen chloride, various nitriles, and aldehydes (4-12). They are suspected of facilitating slow halocarbon formation through hydrolysis and halogen transfer reactions (13). Much of the dissolved organic nitrogen in natural waters and wastewaters consists of amides, which occur in both proteinaceous and nonproteinaceous forms (14-20). The chemical role of amides during water disinfection has never been clearly elucidated. On one hand, various workers have * Corresponding author phone: (973)540-2033; e-mail: [email protected]. Present address: Pharmaceutical Delivery Systems, Parke-Davis, 170 Tabor Rd., Morris Plains, NJ 07950-2536. 3568

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claimed that amides are essentially inert to chlorination in dilute solution (21-23). Based on the work of Margerum et al. (2), rates of amide-N chlorination are expected to be very slow, owing to extremely weak basicity (pKa’s on the order of -1). On the other hand, some workers have presented evidence that amides do undergo chlorination under mild conditions (1, 24, 25). Ayotte and Gray (24) showed that chlorinated N-acetylglycine would oxidize I- at low pH, implying that chlorinated amides are chemically reactive oxidants and thus potential reservoirs of disinfection power. The trans amide bond has been found to dominate in both peptides and proteins as well as low molecular weight amides (26, 27). The trans amide unit in peptides is often described as existing in two resonance structures:

The right-hand structure, containing a C,N double bond, accounts for the short C,N interatomic distance in proteins and peptides (1.33 Å) as well as the planar character of the amide group. In principle, Cl+ could attach to the amide unit either at the N atom or at the O atom. In the first case, Cl would displace the amide H, and electron withdrawal by Cl+ could destabilize the double bond, possibly permitting rotation around the C,N bond. In this paper, we use proton NMR to study chlorination and dechlorination of model amide compounds. In contrast to previous studies, our measurements have been made at environmentally relevant, near-neutral pH. The NMR approach allows us to observe loss of the amide proton during chlorination and its restoration during dechlorination. From evidence that chlorinated amides undergo rotation on the C,N bond, we propose that Cl attaches to the N atom.

Equipment, Materials and Methods Reagents. Peptides (levorotatory) were obtained from Sigma Chemical Co. and used without further purification. All peptides had their terminal N in acetylated form so that all nitrogens would be present as amides. Johnson Matthey, Inc. reagent grade 5% NaOCl was used to make stock solutions of aqueous chlorine. Purified water was obtained from a Milli-Q water filtration system. Dilutions of sulfite were made daily from reagent grade Na2SO3 (J.T. Baker Chem. Co.) and Milli-Q filtered water. Chlorine solutions were standardized by the amperometric titration method (pH 4) with phenylarsine oxide as the titrant (28). Equipment and Methods. A Bruker AMX500 NMR spectrometer was used to monitor the reaction of protected amino acids and peptides with aqueous chlorine. The test compounds were investigated in aqueous solution, containing 10-18% deuterium oxide to provide a frequency lock. To synthesize a chloramide, aqueous NaOCl (pKa HOCl ) 7.5) was added to a buffered amide solution and mixed rapidly. The mixture was then placed in an NMR tube, the D2O was added, and the sample was allowed to react for the required length of time, normally in the dark at ambient temperature. In some experiments, samples were dechlorinated with sulfite after obtaining the spectra of the chlorinated substrate. This was accomplished by adding a stoichiometric excess of sulfite solution to the NMR tube and rapidly mixing by hand. Two different pulse sequences were used: DANTE (delays alternating with nutations for tailored excitation) on echo water suppression and p11 Echo water suppression (29, 30). 10.1021/es980878e CCC: $18.00

 1999 American Chemical Society Published on Web 09/03/1999

These methods are both capable of performing selective excitations in which proton resonances of the analytes can be accurately measured in the presence of water that is more abundant by orders of magnitude. One can not only probe the conformations of different substrates in fairly dilute aqueous solutions but also lower sample concentrations to minimize intermolecular interactions and concentration dependent side reactions. Additionally it is possible to see resonances where the protons are exchanging with the solvent. A drawback of these methods is that they will not reveal resonances in the vicinity of water (4.63 ppm). Our NMR spectra all show a disturbance in the vicinity of 4.63 ppm. This is a vestige of the water suppression technique. The inverted resonances seen downfield from 4.63 are also caused by the water suppression technique. The 1-1 echo which was used for most experiments (30) gave a far better attenuation of the water proton signal when compared to CW presaturation (typically 100 times better). Additionally, this method will not attenuate any exchangeable protons. The DANTE pulse sequence (29) offered similar advantages as well. A selective proton probe was utilized to obtain the water suppressed proton spectra. The 90 degree pulse width used in the 1-1 echo sequence was 10 µs at a power level of 0 dB (utilizing a BSV-10 E-coupler 100 W linear amplifier). All measurements were collected with spectral width, and data size was adjusted to achieve a digital resolution of 0.5 Hz per data point. Typical data size SI is 16K words. Relaxation delay is typically 2-3 s. Temperatures were controlled to within (0.1 C of the desired value with a Bruker temperature controller (Model Eurotherm W1100512). The raw data, free induction decay (FID) was treated with the same exponential weighting function (LB of 0.8 Hz) used to filter out the noise before Fourier Transformation. The value of LB was determined by observing the natural line width of the proton signals from the spectrum without any filter function. Chemical shifts were calibrated against DSS (3-(trimethylsilyl)propanesulfonic acid sodium salt). The external reference DSS was dissolved in D2O and buffer in a separate sealed capillary and did not come in contact with the samples. In the variable temperature studies, calibration was carried out at each temperature with the DSS signal from the external capillary which contained buffer and D2O at the same concentration, temperature, and pH as the sample. For the ambient temperature studies the temperature was actively controlled at 298 K. At 298 K, calibration was accomplished by the external reference DSS control held at the same temperature. At ambient temperature the HDO signal was found to be 4.63 ppm. For subsequent experiments at 298 K the HDO resonance was assigned a value of 4.63 ppm and used to calibrate the spectra. For the ambient temperature experiments the composition of the samples was very similar. Some slight differences in the chemical shifts observed can be attributed to variations in the magnetic susceptibilities caused by small changes in sample pH, ionic strength, reagent concentration, or temperature (when spectra held at different temperatures are compared).

Results N-Acetylglycine. Chlorination of 4.0 mM N-acetylglycine in a 0.10 M phosphate buffer with 5.0 mM HOCl/OCl- at pH 6.6 for 25 h produces intriguing spectral features. The standard with no chlorine (Figure 1A) shows the expected N proton at 7.85 (singlet), the R carbon protons at approximately 3.60 (doublet), the acetyl methyl protons at 1.90 (singlet), and a small unknown peak at 1.80 (singlet). After reaction with HOCl/OCl- for 25 h, two new peaks appear near the methyl group (Figure 1B). These are a broad unknown at 2.05 (singlet) and broad unknown at 2.20 (singlet). The unknown peak at 1.80 (singlet) appears larger. When the

FIGURE 1. A: 4.4 mM N-acetylglycine in 0.10 M phosphate buffer, pH 6.58. A (inset): r carbon resonance from an independent experiment run under similar conditions. B: 4.01 mM N-acetylglycine and 5.3 mM NaOCl after 25 h reaction at ambient temperature; 0.10 M phosphate buffer, pH 6.58. C: Dechlorination of the chlorinated sample with excess SO32-, 15 min reaction time. Conditions: p11 echo water suppression, 256 scans, 298 K. Peak at 4.63 is residual water, which is identified with an asterisk in this and subsequent spectra.

FIGURE 2. A: 5.0 mM N-acetylglycine and 9.0 mM NaOCl in 0.10 M phosphate buffer, pH 6.97, 9 day reaction. B: Reaction of above with excess SO32-; p11 echo water suppression, 256 scans, 298 K. oxidant in the sample is quenched with excess sulfite (Figure 1C), the original spectrum is regenerated, suggesting minimal decomposition of the substrate. Preliminary work involved the chlorination of a number of different substrates followed by a thin-layer assay which could directly detect N-chloro species (25). In the case of a substrate like N-acetylglycine, where only the amide N could react with HOCl/OCl-, it was noted that the reaction product formed slowly on a time scale of hours. Based upon this observation, it is believed that the middle spectrum of Figure 1 represents a reaction which has not gone to completion. The spectrum probably is that of a mixture of unreacted substrate and products. To ascertain whether chlorinated peptides involve formation of Cl-N bonds or Cl-O bonds, a longer (9-day) experiment was done. Deuterium exchange would diminish the amide proton resonance by approximately 18% under these conditions, as the sample contained 18% D2O by weight. Halogenation at the N would further diminish the N-H resonance intensity. Figure 2A shows that the N-bound proton at 7.8 ppm disappears after this lengthy equilibration. Additionally, the R carbon doublets at 3.60 ppm are no longer visible. These resonances have been shifted downfield, nearer to 4.63, where they cannot be observed due to water VOL. 33, NO. 20, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 4. Proposed mechanism of cis-trans isomerism.

FIGURE 3. A: 7.5 mM N-acetylglycine and 11.3 mM NaOCl in 0.20 M phosphate buffer at pH 6.72 and 300 K, 28 h equilibration time. B: 305 K, C: 310 K, D: 320 K Conditions: p11 echo water suppression, 256 scans, DSS used as an external standard. suppression. Halogenation at oxygen and subsequent deprotonation at the amide nitrogen could account for the changes in the amide proton resonance observed but is unlikely given the mild experimental conditions. Additionally, attenuation of the R carbon proton resonances due to Cl-O bond formation would be a longer range interaction than that observed from N-Cl. These observations suggest that the spectrum may be interpreted in terms of N-Cl. The four peak pattern is also visible in the vicinity of 2.0 ppm, although the area ratio of the peaks is somewhat different. Again, dechlorination with excess sulfite regenerates the original spectrum (Figure 2B) with the peak at 1.80 (singlet) appearing larger. At the request of an anonymous reviewer, a control experiment was run under the same experimental conditions in the absence of chlorine or sulfite. The proton bound to the nitrogen of N-acetylglycine was clearly observed after an 11 day equilibration, indicating that deuterium exchange alone cannot account for the complete diminishment of the N-H resonance seen in the preceding experiment. The peak at 1.80 did appear larger after the 11 day equilibration of the control sample when compared to the freshly made solution, suggesting that some degradation of the substrate may be occurring in the control. The control spectra after 11 days was similar to Figure 2B and showed minimal decomposition in the absence of oxidant. The four-resonance pattern seen at approximately 2.0 ppm is investigated as a function of temperature in Figure 3. Some slight changes in the experimental conditions were observed to effect the magnetic susceptibilities and shift the spectrum downfield (to the left) approximately 0.10 ppm. Note that the two broad peaks downfield from the methyl resonance at approximately 2.0 coalesce together over the temperature range of this experiment. At 320 K they have merged into a broad peak centered between the two original peaks (approximately 2.2 ppm). The following model is proposed to explain these observations (Figure 4). The substrate is originally in the thermodynamically stable, trans conformation (B). It slowly reacts with HOCl to form the trans chloramide (A), which is in rapid equilibrium with the cis isomer (C). This rapid equilibrium accounts for the broadness of the chloramide peaks (A and C). The energy barrier between these two conformations is low. In the variable temperature experiment, the cis (C) and trans (A) resonances of the chloramide merge together at higher temperatures as their lifetimes decrease with increasing temperature. At 320 K the cis and trans isomer are seen as a single resonance due to their extremely short 3570

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FIGURE 5. A: N-acetylalanine standard (5.9 mM) in 0.10 M phosphate buffer, pH 6.61. B: 8.4 mM N-acetylalanine with 8.4 mM NaOCl in 0.10 M phosphate buffer, 19 h reaction time. B (inset): Spectrum of side chain CH3 group at 320 K. C: Dechlorination of the sample with excess sulfite; pH after dechlorination 6.56. Conditions: p11 echo water suppression, 256 scans, 298 K. lifetimes on the NMR time scale. The amide nitrogen is weakly basic. Hydrolysis (back reaction) with water will form the unchlorinated substrate but now in two conformations (B and D). This is probably due to return of the structural rigidity of the peptide linkage when the chlorine leaves. The cis form of the unchlorinated substrate would be a thermodynamically unstable form of the molecule. It would isomerize back to the trans form but probably slowly. Resonances seen in Figure 3 at 300 K are assigned as follows: 2.3 ppm (A), 2.15 ppm (C), 2.0 ppm (B), 1.9 ppm (D), 3.75 ppm (R carbon), and 8.0 ppm (nitrogen). It has been suggested that the peak appearing at approximately 1.80 ppm in Figures 1 and 2, which also appeared in Figure 3, and has been assigned structure (D) may be acetate. This material would presumably be present initially as an impurity, with chlorination of the substrate possibly accelerating the formation of acetate via hydrolysis. The size of this small peak increased very slightly during the chlorination/dechlorination process (see Figure 1). This would indicate a minor degradative pathway under the reaction conditions described. N-Acetylalanine. The top spectrum in Figure 5 is unchlorinated N-acetylalanine. The following summarizes the peaks: NH at 7.8 (singlet); acetyl methyl group at 1.9 (singlet); and the side chain methyl at 1.2 (doublet). The resonance of the R carbon proton is obscured by the water suppression technique. The disturbance at 4.63 is also from water suppression. Reaction with HOCl/OCl- (Figure 5B) produces two major new peaks: a broad peak downfield from the side chain resonance at about 1.4 ppm (singlet) and a sharper peak downfield from the acetyl methyl resonance at approximately 2.2 ppm (singlet). The magnitude of the change in chemical shift seen for the acetyl methyl group (∂δ ) 0.3 ppm) corresponds to that observed for the trans isomer of N-chloro-N-acetylglycine. This observation is consistent with

FIGURE 6. Concentration of chlorinated N-acetylalanine formed vs time. Conditions: 0.10 M phosphate buffer, pH 7.02, 298 K, DANTE water suppression, N-acetylalanine 3.8 mM, NaOCl 7.6 mM. The curve is calculated by fitting eq 1 (text) to the data. steric hindrance, which would be expected between the alanine side chain and acetyl methyl group and which would restrict formation of the cis conformation. Dechlorination with excess sulfite (Figure 5C) again regenerates the original spectrum, implying minimal substrate decomposition. Again, the middle spectrum (B) is believed to be a mixture of unreacted substrate and N-chloro-N-acetylalanine. Due to sluggish kinetics of chlorination, the amide proton resonance is visible. A variable temperature experiment was done to better understand the spectral features in Figure 5. The broad peak at 1.40 ppm broadened into a doublet at 320 K, suggesting the presence of two distinct chemical entities which would deshield the side chain methyl group differently (see Figure 5B, inset for spectrum at 320 K). This peak is interpreted as two overlapping resonances (instead of a doublet). This suggests the hybridization of the amide N is closer to sp3 when chlorine is bound. There are two isomers, one with chlorine syn to the alanine side chain and one with chlorine anti to the alanine side chain. The other spectral features were essentially unchanged at 320 K, except for the relative areas of the peaks. Note that the deshielded side chain has grown larger at the elevated temperature, which favors the formation of the chloramide group. The rate of chlorination of N-acetylalanine in 0.1 M phosphate buffer at pH 7.0 was investigated in an 8-day experiment. The concentration of the chlorination product was determined by manually integrating the deshielded acetyl methyl peak (2.2 ppm). The spectrometer was programmed to do one 256 scan experiment each hour overnight and then one experiment each day until the system approached equilibrium. The water signal was suppressed with DANTE excitation (on resonance), which resulted in more accurate integration. The results are plotted in Figure 6. The data were fit by numerically integrating the following differential equation, adjusting kf and kb

δ(Cl-AA) ) kf[HOCl][AA] - kb[Cl-AA] δt

(1)

where kf is a second-order rate constant for the (forward) chlorination reaction, kb is a first-order rate constant for the hydrolysis of the chloramide (back reaction), AA represents N-acetylalanine, and Cl-AA represents its chlorinated form. Consistent with the chlorination of amines in general (2), we assume that the chlorination rate is first order in both peptide and HOCl. The fit yields kf ) 1.58 ( 0.38 × 10-3 M-1 s-1 and

FIGURE 7. A: 4.6 mM N-acetylalanylalanylalanine, 0.20 M phosphate buffer, pH 6.72; B: 5.1 mM N-acetylalanylalanylalanine with an equimolar amount of NaOCl, 22 h equilibration time, 0.20 M phosphate buffer; C: dechlorination of the N-chloropeptide with excess sulfite, the spectra is obtained approximately 15 min after dechlorination; conditions: 298 K, p11 echo water suppression, 256 pulses. kb ) 7.57 ( 1.15 × 10-7 s-1. Other rate laws were tested, but none produced the excellent fit achieved with eq 1. From these rate constants, the equilibrium constant is derived:

K)

kf [Cl-AA] ) ) 2.1 ( 0.6 × 103 kb [HOCl][AA]

(2)

Because we were not able to repeat this lengthy experiment at multiple pH’s and buffer concentrations, these constants must be regarded as conditional constants, valid only for pH 7.0, 0.1 M phosphate, and 298 K. At the end of the 8 day experiment, total oxidant was determined by amperometric titration. It was determined that only 14% of the total oxidant was lost during the lengthy equilibration. As the titration detects both free chlorine and N-Cl bound chlorine, this is additional evidence of chloramide formation where most of the Cl that reacted with the peptide remained an active oxidant. N-Acetylalanylalanylalanine. The protected peptide, Nacetylalanylalanylalanine, was allowed to react with HOCl/ OCl- in 0.2 M phosphate buffer at pH 6.7. The top spectrum of Figure 7 shows the unchlorinated peptide. The peaks are identified as follows: 1.2 ppm - side chain methyl groups; 1.9 ppm - acetyl methyl protons; R carbon protons are not seen due to the water suppression technique. The N-H resonances have been assigned as follows: amide 1, 7.74 ppm; amide 2, 8.16 ppm; amide 3, 8.07 ppm. The middle spectrum (B) illustrates the effects of chlorination with equimolar quantities of peptide and aqueous chlorine. A new peak corresponding to deshielded, side chain methyl groups is visible at 1.4 ppm. There is a new resonance downfield from the methyl on the acetyl group at 2.2 ppm. The magnitude of the chemical shifts observed are consistent with those seen in the N-acetylalanine experiment. These new peaks are consistent with the deshielding effects, which would be expected from a distribution of N-bound Cl along the amide backbone of the peptide, with the reaction conditions forming a mixture of monochlorinated isomers. The reaction shown in Figure 7 illustrates chlorination of the mid amide N (2), but two other isomers could also form with N-bound chlorine at site (1) or (3). The amide proton resonances show an interesting deshielding effect (Figure 7B). Each original resonance is visible, as the reaction had not gone to completion (the chlorine dose here would chlorinate 1/3 of the amide N sites). VOL. 33, NO. 20, 1999 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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To the left of each amide hydrogen resonance is a smaller resonance corresponding to the same amide hydrogen which is deshielded by chlorine on the adjacent amide nitrogen. Farthest downfield (8.3 ppm) is a resonance assigned to amide nitrogen 2. The greater intensity of this deshielded N-H resonance results from two resonances approximately equal in chemical shift, where the mid amide N (2) is deshielded by monochloropeptides with chlorine bound at positions N (1) or N (3). The bottom spectrum (Figure 7C) illustrates regeneration of the original spectrum through dechlorination, with minimal oxidative decomposition.

Discussion There is little information in the scientific literature concerning the rates of chlorination of amides. An uncatalyzed, second-order rate constant of 0.05 M-1 s-1 has been reported for the reaction of N-acetylglycine with HOCl at pH 6 (1). Ayotte and Gray (24) reported that Cl2 was more effective than HOCl when N-acetylglycine was chlorinated. At pH 1.1, an observed rate constant of 0.066 s-1 (second-order rate constant of 0.67 M-1 s-1) was reported for the reaction of Cl2 with N-acetylglycine; HOCl was about 10 times less effective as a chlorinating agent. The rate measurements reported here are significantly slower than these previous studies suggest, implying that low pH conditions used in those studies catalyzed the chlorination reaction. The results described here, of stable N-chloro species showing minimal oxidative decomposition on a time scale of days, is consistent with ref 24, where N-chloro-Nacetylglycine was found to be a stable oxidant species over 48 h at acidic pH. Recently, Hawkins and Davies (31) described rapid oxidant demand when N-methylacetamide was chlorinated near neutral pH in a higher concentration range than reported in this study. The mechanism of oxidant demand responsible for this disparity in results is poorly understood at this time and may possibly be concentration dependent. Schiller et al. described a broad proton resonance at 2.35 ppm observed in the proton NMR spectra of the supernatant from chlorinated porcine cartilage (32). The identity of this species is unknown at this time, though the results here suggest the presence of an N-chloro-N-acetyl functional group. To the extent that N-acetylalanine can be invoked as a model for peptides and proteins in natural waters and wastewaters, our data suggest that chlorinated amides will not be important in disinfection chemistry. Given the equilibrium constant determined for N-acetylalanine, the amide unit could be only 2% chlorinated at an HOCl concentration of 10-5 M (0.7 ppm as Cl2, i.e., representative of disinfection conditions). A more stringent constraint is posed by the rate of formation of Cl-N-acetylalanine. For a constant HOCl concentration of 10-5 M, approximately 1.5 years would be needed to achieve 50% conversion of the amide to its chlorinated form. To the extent that this rate may be representative of amides in general, it would appear that appreciable Cl-amide can form at near-neutral pH only at very high HOCl concentrations. However, if once formed (e.g. in a high-concentration sidestream such as sometimes used in water chlorination), then Cl-peptides would hydrolyze quite slowly. They might persist in solution for a sizable period in the absence of reducing agents. They apparently are reactive to I- on the time scale of a titration and thus might be reactive as well to microbial targets, providing some disinfection capacity. This remains to be determined. The mechanism of cis-trans isomerism observed with N-chloro-N-acetylglycine in this study has not been previously described. It is of interest that the effect was observed at ambient temperature. In biopolymers, the trans peptide linkage is the dominant form, with the cis peptide bond often 3572

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associated with proline (26). Laplanche and Rogers (27) have documented cis-trans isomerism of N-substituted formamides at 35 °C. They determined that the effect was inhibited when the hydrogen of the formyl group was replaced by a bulkier substituent. This is analogous to the observation in this investigation that cis-trans isomerism was inhibited by replacing the R-carbon proton of N-acetylglycine with a methyl group. N-Methyl-N-acetylsarcosinate has also been found to exhibit this type of isomerism but also at elevated temperatures (33).

Acknowledgments This work supported by the Maryland Water Resources Research Center with funds from the U.S. Geological Survey.

Literature Cited (1) Morris, J. C. In Principles and Applications of Water Chemistry; Faust, S. D., Hunter J. V., Eds; John Wiley and Sons: New York, 1967; pp 23-53. (2) Margerum, D. W.; Gray, E. T.; Huffman, R. P. In Organometals and Organometalloids, Occurrence and Fate in the Environment; Bellama, J. M., Brinckman, F. E., Eds; ACS Symposium Series 82; American Chemical Society: Washington, DC, 1978; pp 278291. (3) White, G. C. The Handbook of Chlorination, 3rd ed.; Van Nostrand Reinhold: New York, 1992; pp 889-934. (4) Le Cloirec, C.; Martin, G. In Water Chlorination, Chemistry, Environmental Impact and Health Effects; Lewis Publishers: Chelsea, MI, 1985; pp 821-834. (5) Trehy, M. L.; Yost, R. A.; Miles, C. J. Environ. Sci. Technol. 1986, 20, 117-122. (6) de Leer, E. W. B.; Baggerman, T.; van Schaik, P.; Zuydeweg, C. W. S.; de Galan, L. Environ. Sci. Technol. 1986, 20, 1218-1223. (7) Hirose, Y.; Maeda, N.; Ohya, T.; Nojima, K.; Kanno, S. Chemosphere 1988, 17, 865-873. (8) Young, M. S.; Uden, P. C. Environ. Sci. Technol. 1994, 28, 17551758. (9) Antelo, J. M.; Arce, F.; Paraj, M. J. Phys. Org. Chem. 1996, 9, 447-454. (10) Armesto, X. L.; Canle, L. M.; Garcia, M. V.; Losada, M.; Santaballa, J. A. J. Phys. Org. Chem. 1996, 9, 552-560. (11) Keefe, D. J.; Fox, C.; Conyers, B.; Scully, F. E., Jr. Environ. Sci. Technol. 1997, 31, 1973-1978. (12) Fox, T. C.; Keefe, D. J.; Scully, F. E., Jr.; Laikhter, A. Environ. Sci. Technol. 1997, 31, 1979-1984. (13) Haas, C. N.; Topudurti, K. In Disinfection By-Products in Water Treatment; Minear, R. A., Amy, G. L., Eds.; Lewis Publishers: Boca Raton, Fl, 1996; pp 339-350. (14) Tuschall, J. R., Jr.; Brezonik, P. L. Limnol. Oceanogr. 1980, 25, 495-504. (15) Ram, N. M.; Morris, J. C. Environ. Intl. 1980, 4, 397-405. (16) Parkin, G. F.; McCarty, P. L. Water Res. 1981, 15, 139-149. (17) Scully, F. E.; Howell, G. D.; Kravitz, R.; Jewell, J. T. Environ. Sci. Technol. 1988, 22, 537-542. (18) Scully, F. E.; Hartman, A. C.; Rule, A.; LeBlanc, N. Environ. Sci. Technol. 1996, 30, 1465-1471. (19) McCarthy, M. D.; Hedges, J. I.; Benner, R. Science 1998, 281, 231-234. (20) Frolund, B.; Keiding, K.; Nielson, P. H. Water Sci. Technol. 1994, 29(7), 137-141. (21) Pereira, W. E.; Hoyano, Y.; Summons, R. E.; Bacon, V. A.; Duffield, A. M. Biochim. Biophys.Acta 1973, 313, 170-180. (22) Stelmaszynska, T.; Zgliczynski, J. Eur. J. Biochem. 1978, 92, 301308. (23) Hureiki, L.; Croue, J. P.; Legube, B. Water Res. 1994, 28(12), 2521-2531. (24) Ayotte, R. C.; Gray, E. T. In Water Chlorination: Chemistry, Environmental Impact and Health Effects; Jolley, R. L., Bull, R. J., Davies, W. P., Katz, S., Roberts, M. H., Jr., Jacobs, V. A., Eds.; Lewis Publishers: Chelsea, MI, 1984; Vol. 5, pp 797-806. (25) Jensen, J. S. Chemical Studies to Understand the Dechlorination Process Used at Wastewater Treatment Plants. Ph.D. Dissertation, University of Maryland, College Park, MD, 1997; 145 pp. (26) Wuthrich, K. NMR In Biological Research: Peptides and Proteins; American Elsevier: New York, 1976; pp 184-196.

(27) LaPlanche, L.; Rogers, M. T. J. Am. Chem. Soc. 1964, 86, 337341. (28) Standard Methods For The Examination of Water and Wastewater, 18th ed.; Greenberg, A. E., Clesceri, L. S., Eaton, A. D., Eds.; American Public Health Association: Washington, DC, 1992. (29) Morris, G. A.; Freeman, R. J. Magn. Reson. 1978, 29, 433-462. (30) Sklenar, V.; Bax, A. J. Magn. Reson. 1987, 74, 469-479. (31) Hawkins, C. L.; Davies, M. J. Free Radical Biol. Med. 1998, 24(9), 1396-1410.

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Received for review August 26, 1998. Revised manuscript received June 30, 1999. Accepted July 6, 1999. ES980878E

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