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Schmidt, S. K.; Alexander, M.; Shuler, M. L. J . Theor. Biol.
50-99-7; nystatin, 1400-61-9;amphotericin B, 1397-89-3;ammonium nitrate, 6484-52-2.
1985,114, 1-8.
Yordy, J. R.; Alexander, M. Appl. Environ. Microbiol. 1980,
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39, 559-565.
Jobson, A.; McLaughlin, M.; Cook, F. D.; Westlake, D. W. S. Appl. Microbiol. 1974, 27, 166-171. Law, A. T.; Button, D. K. J. Bacteriol. 1977,129,115-123. Wurtzbaugh, W. A.; Horne, A. J. Can. J. Fish. Aquat. Sei.
1979,37,1211-1216.
Jannasch, H. W. Limnol. Oceanogr. 1967, 12, 264-271. Atlas, R. M.; Bartha, R. Environ. Sei. Technol. 1973, 7,
1983, 40, 1419-1428.
538-541.
MOSS,B. J . Ecol. 1973,61, 193-211. Maloney, T. E.; Miller, W. E.; Shiroyama,T. Spec. Symp.
Lewis, D. L.; Kollig, H. P.; Hodson, R. E. Appl. Environ. Microblol. 1986,51, 598-603. Veldkamp, H.; Jannasch, H. W. J. Appl. Chem. Biotechrwl. 1972,22, 105-123. Subba-Rao,R. V.; Rubin, H. E.; Alexander, M. Appl. Environ. Microbiol. 1982, 43, 1139-1150. Lehtomaki, M.; Niemala, S. Ambio 1975, 4, 126-129. Tagger, S.; Branchi, A.; Julliard, M.; LePetit, J.; ROUX, B. Mar. Biol. (Berlin) 1983, 78, 13-20. Brunner, W.; Sutherland, F. H.; Focht, D. D. J . Environ. Qual. 1985,14, 324-328. Goldstein, R. M.; Mallory, L. M.; Alexander, M. Appl. Environ. Microbiol. 1985, 50, 977-983. Hoben, H. J.; Somasegaran,P. Appl. Enuiron. Microbiol. 1982,44, 1246-1247. Boethling, R. S.;Alexander, M. Environ. Sei. Technol. 1979, 13,989-991. Schmidt, S. K.; Alexander, M. Appl. Environ. Microbiol. 1985,49,822-827.
Am. SOC.Limnol. Oceanogr. 1972,1, 134-140. Harder, W.; Dijkhuizen, L. Annu. Rev. Microbiol. 1983,37, 1-23. Harder, W.; Dijkhuizen, L. Philos. Trans. R. SOC.London, B. 1982,297,459-479. Lyr, H. In Plant Disease; Horsfall, J. G., Cowling, E. B., Eds.; Academic: New York, 1977; Vol. 1, pp 239-261. Wiggins, B. A.; Jones, S. H.; Alexander, M. Appl. Environ. Microbiol. 1987, 53, 791-796. Received for review September 3, 1987. Revised manuscript received March 10,1988. Accepted June 9,1988. This research was supported by funds provided by Public Health Service Training Grant ES-07052 from the Division of Environmental Health Sciences, National Institutes of Health, and by the Army Research Office.
Isolation and Identification of Reaction Products Arising from the Chlorination of Cytosine in Aqueous Solution Gllllan L. Reynolds, Helen1 A. Filaderll, Alun E. McIntyre, Nlgel J. D. Graham, and Roger Perry*
Public Health and Water Resource Engineering Section, Department of Civil Engineering, Imperial College, University of London, London SW7 2BU England
The reaction between free available chlorine and the pyrimidine base cytosine has been studied under controlled conditions of pH and chlorine dose. Reaction mixtures were separated and analyzed by reversed-phase highperformance liquid chromatography. Eluant fractions corresponding to the major UV-absorbing reaction products were further analyzed by field desorption mass spectrometry and nuclear magnetic resonance spectroscopy. The reaction products identified were 1-, 3-, and 5-chlorocytosine and 3,5-dichlorocytosine. 1- and 3chlorocytosine were formed in high yields at both neutral to low and high pH values with a formation minimum at pH 9.0. 5-Chlorocytosine and 3,5-dichlorocytosine were produced in highest yields under alkaline conditions with a progressive decrease in production as pH decreased. The aromatic reaction products identified were formed in highest yields at [chlorine]:[cytosine] molar ratios of 2-3. Nonaromatic reaction products became increasingly predominant with increase in the initial [chlorine]:[cytosine] molar ratio.
Introduction Increased awareness of the possible relationship between human disease and environmental pollutants has resulted in the necessity for the measurement and characterization of both natural and synthetic organic compounds in raw and potable waters. To date, the majority of concern over organic contaminants in potable water has centered on the volatile compounds, particularly trihalomethanes, for which quantitative analytical methodology has been developed (I). This 0013-936X/88/0922-1425$01.50/0
concern has been reflected in the promulgation of water quality standards (2). However, it is now accepted that only 20% of the organic matter present in water is volatile, and interest is currently being directed toward the nonvolatile organic fraction and in particular to the identification of those compounds resulting from the chlorination of raw and waste waters (3). Among some of the more biologically active nonvolatile organic compounds that have been identified in raw and potable waters are the N-heterocyclic purine and pyrimidine bases (41, which are the fundamental building blocks from which nucleic acids are constructed. Early chlorination studies involving purines and pyrimidines were carried out in order to elucidate the mechanisms of microbial and viral inactivation during the disinfection process (5-7). Current interest in the chlorination reaction of these bases follows the display of mutagenic activity (8) and the toxicity of some chlorinated base derivatives (9). General agreement on the relative reactivity of purines and pyrimidines toward chlorination has previously been found: the purine bases adenine, guanine, and xanthine tend to be resistant to attack by aqueous chlorine (IO), while the pyrimidine bases uracil, cytosine, and thymine appear to be significantly more reactive and undergo halogen substitution reactions or ring cleavage (6,7,11-15). However, the substrate and/or disinfectant concentrations used in these studies have generally been far higher than those encountered under typical water-treatment conditions, and the reactions occurring and products formed may differ from those found at lower concentrations (16). In addition, reactions have been monitored by nonspecific
0 1988 American Chemical Society
Environ. Sci. Technol., Vol. 22, No. 12, 1988 1425
techniques (e.g., a change in UV absorption) or more specific techniques (e.g., gas chromatography and gas chromatography-mass spectrometry) inappropriate for the characterization of heterocyclic compounds of low volatility. To obtain a better understanding of the reaction of pyrimidines during disinfection by chlorine, more specific and reliable analytical techniques able to separate and identify individual compounds of concern have been applied. These techniques included high-performanceliquid chromatography (HPLC), field desorption low-resolution mass spectrometry (FD/MS), proton nuclear magnetic resonance spectroscopy (NMR) and thin-layer chromatography (TLC).
Experimental Section Procedure. Cytosine chlorinations were carried out in glass reaction vessels incubated in a darkened recirculating water bath maintained at 10 f 0.5 “C. The chlorinating agent, 1M sodium hypochlorite in 0.1 M sodium hydroxide solution, was added dropwise to the buffered cytosine solution (50 mg L-l, pH 7) with continuous stirring to give an initial [free available chlorine (FAG)]:[cytosine] molar ratio of 3. Samples were withdrawn at intervals for determination of free and combined chlorine (17)and for separation and fractionation by HPLC. Excess free chlorine was removed from reaction mixture samples prior to HPLC analysis by use of Sep-Pak C18 cartridges [Waters Associates, Millipore (UK) Ltd.] as follows: The cartridge was wetted with l-mL of methanol. A l-mL sample was applied to the cartridge. Inorganics (including free chlorine) were eluted with 0.5 mL of water. Organic compounds were eluted with 1mL of methanol. The methanol eluate was applied to the HPLC column. The major UV-absorbing HPLC eluant fractions were analyzed by field desorption mass spectrometry (FD/MS), nuclear magnetic resonance spectroscopy (NMR), and thin-layer chromatography (TLC) for cytosine chlorination product identification. Further chlorination reactions were carried out at a cytosine concentration of 50 mg L-l and at a constant pH (7.0) with variation in the initial [FAC]:[cytosine] molar ratios (1,2,3, 5 , 7 , 10) and at a constant [FAC]:[cytosine] molar ratio (3) with variation in the pH of the reaction mixture (6.0, 7.0, 8.0, 9.0, 10.0, 11.0). Combined chlorine residual and HPLC analyses of reaction mixture subsamples were carried out at preselected time intervals. Identification Techniques. HPLC was carried out on a Waters 6000A HPLC system equipped with an automated gradient elution controller and a Lambda-Max (Model 480) UV absorption detector operated at 280-nm wavelength. The reversed-phasecolumn (100 mm X 8 mm i.d.; Radial Pak b-Bondapack CI8,particle size 10 pm) was operated in a radial compression module (Waters z-module) with a linear solvent gradient from 0% methanol (100% water) to 100% methanol (0% water) over 30 min at a flow rate of 2.0 mL min-’. Field desorption low-resolution mass spectrometry (FD/MS) was performed on a JEOL DS 300 double-focusing mass spectrometer equipped with a combination FD/FI/EI (field desorption/field ionization/electron impact) ion source and an emitter anode current programmer. After the ion source was tuned and the magnetic field calibrated with perfluorokerosene in the E1 mode, the FD/FI mode was selected and the source was retuned with acetone. Samples (in methanol) were applied by microsyringe to the emitter anode which consisted of 10-pm tungsten wire activated with carbon needles as described by Beckey et al. (18). Low-resolution (-800 m/Am) mass 1426
Mviron. Sci. Technol., Vol. 22, No. 12, 1988
Time
min
Figure 1. Cytosine consumption and reaction product formation over 250-min reaction period (pH 7.0, [FAC]:[cytosine] molar ratio = 3.0).
spectra were obtained by using a 3.5-kV accelerating voltage and -7.5-kV cathode voltage. The emitter current was programmed to increase linearly from 0 to 30 mA at 1 mA min-l. NMR spectra were obtained on a Bruker 250-MHz instrument with hexadeuteriodimethylsulfoxide (DMSO-d6) as solvent and trimethylsilane (TMS) as an internal reference standard. Thin-layer chromatography of HPLC eluant fractions was carried out on silica gel TLC plates (200 X 200 mm, 0.25-mm silica gel G) using a mobile phase consisting of chloroform/methanol/ammonia (90/10/ 1v/v). Developed plates were viewed under UV light (254 nm) to detect UV-absorbing sample components. Chloramines were visualized by spraying with 2% potassium iodide in 5% acetic acid solution, followed by 0.5% aqueous starch solution.
Results Identification of Cytosine Chlorination Products. The results for the preliminary chlorination run, carried out at pH 7.0 with a [FAC]:[cytosine] molar ratio of 3 are presented in Figure 1 where concentrations of cytosine, reaction products, and combined residual chlorine are plotted against time over the course of the experiment. Since reference standards for chlorinated cytosine derivatives were not available, concentrations of these compounds were calculated in terms of “cytosine equivalents”. The cytosine concentration declined rapidly over the first hour of the reaction while the corresponding concentration of combined chlorine increased, reaching a plateau after 100 min. Concentrations of the two major unidentified reaction products gradually increased throughout the 250-min chlorination period. This general pattern was apparent in subsequent chlorination runs. A sample HPLC chromatogram is presented in Figure 2. Five major UV-absorbing chromatographic peaks were apparent. Eluant fractions corresponding to peaks 2-5 were collected and subjected to FD/MS, NMR, and TLC analyses for identification, Preliminary FD/MS analysis of the fraction corresponding to peak 1suggests it arose from inorganic substances. No further analysis was carried out on this fraction. HPLC retention time data suggested that peak 2 corresponded to unreacted cytosine, and FD/MS analysis gave a mass spectrum with a base peak of m/z 111and a weaker ion at m / z 112. Comparison of the spectrum with that of a standard cytosine solution indicated that the FD mass spectrum corresponded to unreacted cytosine. The mass spectrum of the third fraction displayed parent ions at m / z 145 and 147 indicating the presence of M + [36C1]m~n~chlor~cyto~ine and the 37Clisotope, respectively. A similar mass spectrum was obtained for the fourth peak fraction, with a base peak at m / z 145, indicative of a monochlorocytosinecompound. The FD mass
0
dl 1v
V
Figure 5. Probable structures of the monochlorocytosineidentified in the peak 4 fraction (1- and/or 3-chlorocytoslne).
VI
1
Figure 8. Probable structure of the dichlorocytosine identified in the peak 5 fraction (3,5-dichlorocytosine). A
&
14 16
Time (min)
1 0
+ FAC
F A C Cytosine FAC C y t o s i n e
2 0
Cytoslnr
3 0
x FAC
Cvtosine
5 0
0
Figure 2. HPLC chromatogram of cytosine reaction mixture (at 250 min).
II
I
Figure 3. Probable structures of cytosine tautomers identified in the peak 2 fractlon.
I ;+
HOAN 111
Figure 4. Probable structure of the monochlorocytosine identified in the peak 3 fraction.
spectrum of the fifth peak fraction exhibited a base peak at m / z 179, among an isotopic ion cluster characteristic of a dichloro compound, with intense ions corresponding to 36Cl[M]" and 37Cl[MI'+. The ion cluster in the m / z 200-210 region of the spectrum corresponds to the %C1[M Na+]+and %1 [M Na+]+species. This compound was therefore tentatively identified as a dichlorocytosine. Proton NMR spectroscopy was utilized for further analysis of the four HPLC fractions in order to enable structural confirmation of the compounds identified by FD/MS. The NMR spectrum of the second fraction, tentatively identified as unreacted cytosine by FD/MS, exhibited two doublet peaks at 6 5.55 and 7.3, which correspond to the 5- and 6-position protons of cytosine, a wide doublet at 6 7.0, which corresponds to the NH2 group, and a wide singlet peak at 6 10.3, corresponding to the OH group of the enolic form (I) of the two cytosine tautometers (I, 11) illustrated in Figure 3. NMR analysis of the third fraction previously identified as a monochlorocytosine,gave one doublet at 6 7.7, which corresponds to the 6-position proton, and a wide singlet at 6 10.3, which corresponds to the OH group. Distribution of the proton, amino, and hydroxyl groups in such positions indicates thia reaction product to be 5-chlorocytosine (Figure 4). The fourth peak fraction displayed two pairs of doublets at 6 5.55, 5.95, 6.94, and 7.21 likely to be attributable to the 5- and 6-position protons of the monochlorocytosines (IV, V) illustrated in Figure 5. The l-chlorine, adjacent
+
+
Time, min
Figure 7. Cytosine consumption over a range of chlorine doses (pH 7.0).
to the 5- and 6-positions protons, causes large shifts at 6 5.95 and 6.94, while the 3-chlorine, being far from the 5and 6-positions, causes no shifts and the two doublets appear at 6 5.55 and 7.21. The two wide peaks at 6 7.0 and 10.3 correspond to the NH2 and OH groups, respectively. NMR analysis of the fifth fraction, tentatively assigned as dichlorocytosine, produced a spectrum with wide peaks at 6 7.28. This reaction product is most likely to be 3,5dichlorocytosine (Figure 6); since no significant chemical shift of the 6-proton signal occurs, the second chlorine is unlikely to be in the N1 position. At pH 7 and a [FAC]:[cytosine] molar ratio of 3, chlorination of the cytosine 4-amino group did not appear to have taken place. The major products of chlorination appeared to be 1- and 3-chlorocytosine, 3,5-dichlorocytosine, and 5-chlorocytosine. In addition, the formation of a nonaromatic reaction product was apparent from the presence of a non-UV-absorbing compound following TLC analysis of reaction mixture samples. This compound was not identified. The absence of a chlorinated 4-amino derivative conflicts with the results of Patton et al. (7) who reported a 76% yield of 4-chlorocytosine following the reaction of cytosine with 1equiv of HOCl. However, in this study and that of Patton, the NMR data (and melting point and UV spectral data in the latter study) are insufficient to totally exclude the formation of either 4chlorocytosine or other monochlorinated cytosine derivatives, respectively. Effect of Chlorine Dose upon Cytosine Chlorination. Cytosine consumption at varying [FAC]: [cytosine] molar ratios (pH 7.0) is shown in Figure 7. Consumption increased with chlorine ratio up to 51, whereafter all the cytosine present had reacted. The pattern of formation of the chlorinated cytosine derivatives previously identified (as l-chlorocytosine, 3-chlorocytosine, and 3,5-dichlorocytosine) and the combined residual chlorine at varying [FAC]:[cytosine] molar ratios is depicted in Figure 8. Environ. Sci. Technoi., Voi. 22, No. 12, 1988 1427
.
IA
Cytosine + I N a n d l o r 3N-Chlorocytosine
80
x 0
3 , 5-Dichlorocytosine Combined residual c h l o r i n e
90 ; 80
;
170:
70
R"
100
L
60 0
FAC
Cytosine molar r a t i o
Flgure 8. Variation in cytosine consumption and reaction product identity with chlorine dose (pH 7.0).
pH OpH t p H XpH A pH
6 0 7 0 8 0
9 0 10 0
m - - - - T A = +
-
Y ' e ; 100
200
150
d 250
T i m e , min
Figure 9. Cytosine consumption at different pHs ([FAC]: [cytosine] = 3.0). Cytosine
+ 1N andlor x 3,
3N-Chlorocyiosine
5. Dichlorocytosine
0 5 - Chlorocytostne o Combined residual chlorine
PH
Flgure 10. variation in reaction product identity with pH ([FAC]:[cytosine] = 3.0).
5-Chlorocytosine was not detected, although it had been identified in preliminary experiments at pH 7.0 and at a [FAC]:[cytosine] molar ratio of 3 (see previous section). Formation of the N-chlorocytosines reached a maximum at ratios of 2:l and 3:1, and they were not detected at a ratio of 10:l. Peak formation of the reaction product identified as 3,5-dichlorocytosine occurred at a [FAC]: [cytosine] ratio of 3. Higher ratios brought about more rapid initial formation of 3,5-dichlorocytosine; however, gradual decomposition of this compound occurred over the 250-min reaction period.
Combined residual chlorine concentrations increased progressively with increase in the [FAC]:[cytosine] molar ratios. As levels of combined residual chlorine increased the total concentration of identified reaction products tended to decrease (at least at ratios of [FAC]:[cytosine] greater than 3) until at a ratio of 10, when maximum combined residual chlorine levels were recorded, no Nchlorocytosines or 3,5-dichlorocytosine was detected. HPLC and TLC analyses revealed the absence of aromatic, UV-absorbing compounds in the 1O:l [FAC]:[cytosine] molar ratio reaction mixture at the end of the 250-min reaction period. Effect of pH on Cytosine Chlorination. At an initial [FAC]:[substrate] molar ratio of 3:1, cytosine consumption varied with the pH of the reaction mixture in the manner shown in Figures 9 and 10. Maximum consumption was evident at pH 6.0. Consumption declined to a minimum at pH 9.0 and subsequently increased with increasing pH (Figure 10). Formation of the chlorinated cytosine derivatives, identified as l-chlorocytosine, 3-chlorocytosine, 5-chlorocytosine, and 3,5-dichlorocytosine, and combined residual chlorine at the different pHs is also depicted in Figure 10. Concentrations of 5-chlorocytosine were consistently lower than those of the other derivatives determined. Maximum 5-chlorocytosineformation occurred at pH 11.0, Levels decreased progressively with decrease in pH until little or no 5-chlorocytosine was detected in reaction mixtures buffered at pHs below 9.0. A similar pattern of variation in the level of the compound identified as 3,5dichlorocytosine was apparent: Minimum levels were found at the lower pHs (6.0-7.0); maximum levels were found in reaction mixtures buffered at pHs 10.0-11.0. Formation of the N1-and/or Ns-chlorinated derivatives followed an inverse trend to cytosine consumption: highest yields were recorded at pH 6.0; yields decreased with increasing pH reaching a minimum at pH 9. Thereafter levels increased, reaching a value approximately 60% of that recorded at pH 6, as the pH of the reaction mixture increased to pH 11. The pattern of variation with pH in substrate consumption and reaction product formation is consistent with the inverse relationship between the reactivity of the substrate and of the chlorination agent on passing from low to high pH (Figure 11). At low pH, protonation of hypochlorite and cytosine will lead to formation of the powerful chlorination agent hypochlorous acid while susceptibility of cytosine to electrophilic attack, particularly following protonation of the 4-amino group, will be low. As the pH of the reaction mixture rises, the dissociation favors the hypochlorite ion and chlorination efficiency will be poor. Conversely, dissociation of the cytosine 2-hydroxy group at high pH will create a highly nucleophilic site activating the substrate molecule toward chlorination via
HIGH pH
Figure 11. Dissociation of (a) hypochlorous acid and (b) cytosine (79, 20). 1428
Environ. Sci. Technoi., Voi. 22, No. 12, 1988
LOW pH
electrophilic substitution. At pH 9.0, the pH of minimum cytosine consumption, both hypochlorous acid and activated cytosine molecules will be scarce. Little reaction would therefore be anticipated at this pH. Summary
1-and 3-chlorocytosine and 3,5-dichlorocytosine were identified as major producta of reaction of the chlorination of cytosine. Smaller quantities of 5-chlorocytosine were formed. Chlorination of the exocyclic amino group did not appear to have taken place. The formation of nonaromatic reaction products was apparent, and formation increased with increase in the initial [FAC]:[cytosine] ratio. pH significantly influenced the rate of cytosine consumption. The variation in consumption with pH is most likely to be due to the abundance of HOC1 at neutral to low pH and activation of the substrate molecule by dissociation of the 2-hydroxy group at high pH.
(6)
Hayatsu,H.; Pan, S.-K.; Ukita, T.Chem. Pharm. Bull. 1971,
(7)
Patton, W.; Bacon, V.; Duffield,A. M.; Halpern,B.; Hoyano, Y.; Pereira, W.; Lederberg, J. Biochem. Biophys. Res.
19, 2189-2192.
Commun. 1972,48,880-884. (8) Cumming, R. B. In Proceedings of the First Conference on Water ChlorinationEnvironmental Impact and Health Effects,Ann Arbor Science: Ann Arbor, MI, 1975; Vol. 1, (9)
pp 247-258. Gehrrs, C. W.; Eyman,L. D.; Jolley, R. L.; Thomson, J. E.
Nature (London) 1974,249, 675-676. Hoyano, Y.; Bacon, V.; Summons, R. E.; Pereira, W. E.; Halpern, N.; Duffield, A. M. Biochem. Biophys. Res. Commun. 1973,53, 1195-1199. (11) Gould, J. P. Abstracts of Papers, 175th National Meeting (10)
of the American Chemical Society,Anaheim, CA; American Chemical Society: Washington, DC, 1978; ENVR 049. (12) Gould, J. P.; Hay, T. R. Water Sci. Technol. 1982, 14, 629-640. (13)
Gould, J. P.; Richards,J. T.; Miles, M. G. Water Res. 1984,
Acknowledgments
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Gould,J. P.; Richards,J. T.; Miles, M. G. Water Res. 1984,
We acknowledge the advice of M. Fielding and C. D. Watts of the U.K. Water Research Centre. Registry No. Cytosine,71-30-7; 1-chlorocytosine,115651-20-2; 3-chlorocytosine, 115651-21-3; 5-chlorocytosine,2347-43-5; 3,5dichlorocytosine, 115651-22-4.
(15)
Poncir, J.; Martin, G. Enuiron. Technol. Lett. 1986, 7,
Literature Cited (1) Rook, J. J. Water Treat. Exam. 1972,21, 259-272. (2) World Health Organization, Task Group on Guidelines for Drinking Water Quality-Quantification of Selected Health-Related Compounds, Ottawa, Canada, 18-25 Nov 1980, ICP/RCE 209(3), 2917 K, Document A, Second Draft, World Health Organization Regional Office for Europe, Copenhagen, Denmark, 1987; 20pp. (3) Crathorne, B.; Fielding, M.; Steel, C. P.; Watts, C. D. Enuiron. Sci. Technol. 1984, 18, 797-802. (4)
18,205-212. 18,991-999. 177-192.
Fielding, M.; Haley, J.; Watts, C. D.; Corless, C.; Graham, N.; Perry, R. The Effect of Chlorine and Ozone on Organic Compounds in Water-A Literature Reuiew;WRC report, PRD 1217-M, Water Research Centre: Marlow, Bucks, England, 1987. (17) Government of Great Britain, Standing Committee of Analysts Chemical Disinfecting Agents in Water and Effluents,and Chlorine Demand 1980; HMSO: London, UK, 1980. (18) Beckey, H.; Hilt, E.; Schulten, H. J. Phys. E. 1973, 6, (16)
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Levene, P. A.; Bass, L. W.; Simms,H. S. J.Biol. Chem. 1926,
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1979. (5) Prat, R.; Nofre, U.; Cier, A. Ann. Inst. Pasteur, Paris 1968, 114, 595-607.
Received for reuiew October 15,1987. Accepted May 4,1988. We acknowledge the financial support prouided by the U.K. Water Research Centre.
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