Paramagnetic Relaxation of Atrazine Solubilized by Humic Micellar

by concentrated humic acid solutions occupies a domain accessible only to .... used a standard EPR cavity (Bruker ER4102 ST-0). Source of Atrazine and...
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Environ. Sci. Technol. 1997, 31, 3204-3208

Paramagnetic Relaxation of Atrazine Solubilized by Humic Micellar Solutions YAO-YING CHIEN, EUN-GYEONG KIM, AND WILLIAM F. BLEAM* Department of Soil Science, University of WisconsinsMadison, Madison, Wisconsin 53706-1299

This study examines a central tenet of the “membranemicelle” model of humic substancessthat humic molecules form “micelle-like” aggregates with hydrophobic interiors into which nonpolar organic compounds partition. We solubilized atrazine, labeled by a trifluoromethyl group on the ethylamino side chain, in aqueous solutions containing 10% humic acid by weight and observed the F-19 nuclear magnetic resonance (NMR) relaxation of atrazine induced by paramagnetic probes added to the humic solution. One paramagnetic probe, Gd‚EDTA anion, remains in aqueous solution while the other, TEMPO (2,2,6,6tetramethyl-1-piperidinyloxy), is sorbed by humic micelles. The hydrophilic anionic paramagnetic probe induces virtually no paramagnetic relaxation in atrazine solubilized by 10% aqueous humic acid solutions while the hydrophobic neutral paramagnetic probe causes rapid paramagnetic relaxation. These results show that atrazine solubilized by concentrated humic acid solutions occupies a domain accessible only to neutral hydrophobic molecules and, hence, confirms the existence of hydrophobic domains. We suggest that atrazine resides in the hydrophobic interior of humic acid micelles.

Introduction Hydrophobic Organics and Humic Substances. Atrazine, being a photosynthetic inhibitor, is one of the most widely applied herbicides in agriculture. Its recalcitrance toward either chemical or biological degradation and extensive use in agriculture has led to its accumulation in the environment. Detection of atrazine and its metabolites in groundwater has led to moratoria on its use in regions with sandy soils or shallow water tables. Humic substances, more than any other component in soils or sediments, retain atrazine and other xenobiotic organic compounds against leaching by percolating water (1, 2). Lambert (1, 3, 4) described sorption as a partitioning from aqueous solution "into" humic substances. The sorption affinity of neutral organic compounds by humic substances strongly correlates with the hydrophobic character of the organic sorbate (3-8). According to Karickhoff et al. (6, 7), the sorption of hydrophobic organic compounds by organic matter involves “predominantly van der Waals interactions”. Expressions such as hydrophobic (6) or nonpolar (9) characterize humic sorption domains. Molecular Aggregation and Solubilization. The membrane-micelle model (10) describes humic substances as micellar aggregates with hydrophilic exterior surfaces and * To whom correspondence should be addressed; telephone: 806/ 262-9956; fax: 608/265-2595; e-mail: [email protected].

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predominantly hydrophobic interiors. This model explains the sorption and retention of hydrophobic organic compounds as a partitioning into the hydrophobic interior of humic molecular aggregates. Wershaw (11) observed the apparent solubility enhancement of DDT in concentrated humic acid solutions that implicated the formation of humic acid micelles. Evidence for molecular aggregation in humic solutions comes from light-scattering (12), small-angle neutron-scattering (13), and X-ray scattering (14). Numerous studies (15-25) confirm the surface activity of soil, aquatic, and marine humic and fulvic acids. Shinozuka and Lee (24) and Guetzloff and Rice (25) corroborate the solubility enhancement of hydrophobic organic compounds in concentrated humic acid solutions. This behavior parallels that of synthetic surfactants (25, 26) whose capacity to solubilize hydrophobic organic compounds increases significantly above the critical micelle concentration (cmc). Early studies of solubility enhancement by dissolved humic substances (5, 27) found little effect “because of the relatively [small] organic environment and the normally more polar nature of the dissolved organic matter” (5). The reason for limited solubility enhancement became clearer when later studies (24, 25) showed that little or no solubilization occurs below the humic acid cmc. Experimental values for humic acid cmc range from 0.05% to 3% (11, 16, 17, 21-23, 25), much more concentrated than the solutions used by Chiou and co-workers (5, 27) or by von Wandruszka and co-workers (28, 29). Paramagnetic Relaxation Studies. Nuclear dipoles absorb radiofrequency radiation and enter an excited spin state during an NMR experiment, returning to the ground state through numerous relaxation mechanisms. One such pathway involves the interaction between electron and nuclear dipoles that influences both spin-lattice (T1) and spin-spin (T2) relaxation rates (30, 31). Paramagnetic relaxation is very efficient because the electron magnetic moment is much larger than nuclear magnetic moments and result in longrange interactions. Usually, paramagnetic interactions shorten T2 more than T1, though this depends on the magnitude of the electron-nuclear hyperfine coupling constant. The loss of transverse magnetization through spin-spin relaxation shortens the free induction decay (FID) collected in Fourier transform NMR experiments. Line widths in the frequency domain spectrum vary inversely with the duration of the FID. Hence, a direct correlation exists between NMR line widths and the rate of spin-spin relaxation. Paramagnetic relaxation studies of micellar solutions reveal the sorption of hydrophobic paramagnetic probes (32), partitioning of diamagnetic sorbates into micelles (33, 34), counter-ion binding to micelles (35), and location of diamagnetic sorbates within micelles (36). Probes suitable of hydrophobic domains must be water insoluble, while probes of hydrophilic domains must be water soluble and unable to bind to the micellar surface. Electron paramagnetic resonance (EPR) spectroscopy provides an additional means of studying micellization and micellar solubilization. The EPR spectra of nitroxide radicals exhibit hyperfine components whose hyperfine splitting constant (hfsc) correlate with solvent polarity (37). An abrupt change in the nitroxide hfsc occurs at the cmc in aqueous surfactant solutions because nitroxide molecules partition into newly formed micelles (38) where the polarity is lower than the surrounding aqueous solution. The capacity to monitor solvent polarity is essential because not all neutral organic compounds penetrate the micellar core. Chung et al. (39) proposes that phenol buries its aromatic ring in the

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hydrophobic interior while inserting the polar OH group into hydrophilic region at the micelle surface. The more polar hydroquinone, on the other hand, apparently adsorbs entirely at the micellar surface (39). In this paper, we report the differential paramagnetic F-19 NMR line broadening of atrazine induced by hydrophilic and hydrophobic paramagnetic probes. These experiments test the hypothesis that humic acid molecules aggregate in concentrated solutions to form micelles with hydrophobic interiors able to solubilize neutral organic compounds such as atrazine. The humic acid solutions used in this study, though they achieve a 100-fold atrazine solubility enhancement, are far more concentrated than natural humic solutions (5, 27-29). The solubility-enhanced atrazine concentration, however, permits routine NMR observation of atrazine in aqueous humic acid solutions.

Materials and Methods NMR and EPR Instrumentation. The F-19 NMR spectra were collected at the National Magnetic Resonance Facility at Madison (NMRFAM) on a Bruker AM-400 (9.6 T) spectrometer operating at 376.48 MHz. Following a single 5.0-µs pulse, we collected 8 K data points, without proton decoupling, and zero-filled to 16 K before Fourier transformation. Unless otherwise noted, all spectra represent 256 scans separated by 1-s relaxation delay intervals. A standard Bruker heating coil maintained a constant 300 K temperature. All humic acid solutions contained 0.02% (trifluoromethyl)benzene as an internal F-19 reference and 10% D2O for a lock signal. EPR spectra were collected on a Bruker ESP-300E EPR spectrometer. The high dielectric constant of aqueous SDS and humic acid solutions required use of a special sample cell and cavity (Bruker ER4103 TM). For organic solvents, we used a standard EPR cavity (Bruker ER4102 ST-0). Source of Atrazine and Other Compounds. The atrazine (2-chloro-4-trifluoroethylamino-6-isopropylamine-s-triazine) was used as supplied by Ciba Corporation, though purity was verified by thin-layer chromatography. All other chemicals used in this study were reagent-grade and used as supplied. The water was deionized distilled water. We prepared the EDTA (ethylenediaminetetraacetate) complex of Gd(III) as described by Luck and Falke (40), using fresh Gd‚EDTA solutions for each paramagnetic relaxation study. If Gd‚EDTA solutions are stored overnight, crystals begin to form, and the pH of the solution drops below 7.1. We also prepared fresh aqueous solutions of TEMPO (2,2,6,6tetramethyl-1-piperidinyloxy) on the day of the NMR study. Gentle heating of water containing crystalline TEMPO to just below the melting point (36-38 °C) promoted dissolution. Extraction and Preparation of Soil Humic Acids. We extracted humic acid from an alluvial soil (Sparta sand: mesic, uncoated Typic Quartzipsamment) from the lower Wisconsin River valley near Arena, WI. The extraction method followed the procedure recommended by the International Humic Substances Society (41). Rather than using silver nitrate to detect the removal of excess chloride ions, we dialyzed until the conductivity of the external solution reached a plateau at ≈4 µS. We stored the humic acid fraction as a freeze-dried powder. The carbon content and the ash content of the humic acid fraction were 51.3% and 0.65% by weight, respectively. The chemical analysis of the ash appears in Table 1. Iron dominates the inorganic paramagnetics, representing 94% of the total mass of paramagnetic elements. Given the iron content of the ash and the ash content of the humic acid, a 10% humic acid solution would contain 10 ng Fe/mL (equivalent to 10-9 mol of electron spins/mL). We also measured the organic free-radical content of the freeze-dried humic acid by integrating the EPR signal (Figure 1), comparing the integrated peak area to a standard free radical DPPH (1,1-diphenyl-2-picrylhydrazyl). We estimate

TABLE 1. Chemical Content of Residual Ash from Alkaline Sparta Humic Acid Extracta element

ash content (ng g-1)

Fe Cu Cr Ni

151.8 5.8 4.5 2.3

P Ca Na Zn

126.4 39.6 14.3 6.9

element

ash content (ng g-1)

Paramaganetic Co Mn V

2.1 0.7 0.2

Diamaganetic Al Cd Pb

6.8 1.0 0.2

a Concentrations refer to the mass of the element relative to the dry weight of humic acid. Silicon was not measured and probably accounts for much of the remaining ash content.

FIGURE 1. Electorn paramagnetic resonance spectrum of organic free-radicals in freeze-dried Sparta humic acid extract. an organic free-radical content of 1.32 × 10-6 mol of electron spins per gram of humic acid. Newman and Tate (42) found no significant paramagnetic line broadening in C-13 NMR spectra of humic acid solutions containing 2 mg of Fe/mL (equivalent to 10-4 mol of electron spins/mL). We anticipate no significant line broadening arising from paramagnetic impurities, transition metals, or organic radicals in our humic acid samples. Furthermore, since our experimental design measures the differential relaxation rate resulting from added paramagnetic probe, the paramagnetic impurity content of our humic acid would not influence our ultimate results. Preparation of Humic Acid Solutions and Sorption of Atrazine. We dissolved humic acid in alkaline solutions, adjusting the pH to 11.8 with 1 M NaOH. We found that solutions containing 10% humic acid by weight dissolved sufficient atrazine to collect high-resolution F-19 NMR spectra. Atrazine is a solid at 300 K and was supplied as a fine powder. We mixed the desired amount of solid atrazine with freeze-dried humic acid before dissolving in alkaline water. Atrazine solubiliztion in these humic acid solutions typically required stirring overnight. Atrazine sorption experiments were carried out in dialysis cells (molecular mass cutoff ) 1000 Da) with the aqueous humic acid solution confined to one cell and atrazine confined in the other. The atrazine cell contained an initial atrazine concentration of 28 ppm and a C-14 activity of 0.45 nCi/mL. We sampled the atrazine cell hourly to follow the changing atrazine concentration as it diffused across the dialysis membrane into the humic acid cell. We measured the equilibrium atrazine C-14 activity by scintillation counting (scintillation cocktail: Bio-Safe II). We measured and transferred aqueous solutions containing paramagnetic probes (Gd‚EDTA or TEMPO) in a 10-µL

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FIGURE 2. Atrazine solubility enhancement in aqueous solutions containing varying amounts of Sparta humic acid extract. Solubility enhancement of benzo[a]pyrene from Shinozuka and Lee (22) also appear. syringe, mixing these with the micellar solutions directly in 5-mm tubes for NMR studies.

Results Solubility Enhancement. The effect of varying humic acid concentrations on apparent atrazine solubility appears in Figure 2. Varying the pH from 5 to 7 had little effect on atrazine solubility enhancement by the soil humic acid we used. Figure 2 also shows data from Shinozuka and Lee (24) for the solubility enhancement of benzo[a]pyrene in marine humic acid solutions. Although we did not measure the surface tension of these humic acid solutions, we attribute the inflection in the solubility enhancement curve of Figure 2 to the humic acid cmc (24, 25). The sorption capacity, or apparent solubility enhancement, we measured is orders of magnitude higher than that reported in any previous study of atrazine sorption by humic substances (43-52). The conditions of our solubilization experiments differ significantly from those previous studies because our humic acid solutions were far more concentrated and the humic acids were not flocculated. Paramagnetic Relaxation. We measured the relaxation rates of atrazine in concentrated humic acid solutions after adding paramagnetic probes to test our hypothesis that humic micelles contain a hydrophobic interior into which neutral organic compounds partition during solubilization. Our basic assumption is that hydrophobic domains in humic acid solutions exist only when humic acid molecules aggregate. The F-19 NMR spectrum of atrazine in a 10% humic acid solution appears in Figure 3A. Chemical shifts are expresses in parts per million (ppm) relative to the external reference, (trifluoromethyl)benzene. A single atrazine resonance appears in Figure 3A at -9.95 ppm. We attribute the broad shoulder at -9.8 ppm, near the base of the -9.95 ppm atrazine resonance, to other, less stable, atrazine conformers (5356). We confirm our assignment of the -9.8 ppm shoulder by adding increasing amounts of DMSO (dimethylsulfoxide) to the humic acid solution. This progressively resolves the conformers hidden in the -9.8 ppm shoulder (Figure 3B). We selected Gd‚EDTA anion as our hydrophilic paramagnetic probe. The EDTA precludes complexation of the Gd(III) by the humic substances. The most likely location of the Gd‚EDTA anion is in the outer part of the diffuse double layer surrounding negatively-charge humic acid micelles. Our hydrophobic paramagnetic probe is the stable free radical TEMPO. TEMPO has several advantages as a paramagnetic probe: it is a neutral organic compound with low water solubility and its EPR hfsc is sensitive to solvent polarity.

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FIGURE 3. F-19 NMR spectrum of atrazine humic micellar solutions at pH 11.8: (A) 14 mM atrazine dissolved 10% humic acid solution, and (B) 14 mM atrazine dissolved 10% humic acid solution containing 52% DMSO. Internal refererence is 0.02% v/v (trifluoromethyl)benzene in 10% D2O.

TABLE 2. Effect of Solvent Polarity on Electron Paramagnetic Resonance Hyperfine Splitting Constants (hfsc) for TEMPO solvent

hyperfine splitting constant (G)

n-octane benzene n-decane 2-octanone pyridine

15.179 15.321 15.358 15.633 15.708

solvent

hyperfine splitting constant (G)

1-octanol 2-propanol acetic acid water

16.094 16.158 16.609 17.209

Table 2 lists the EPR hfsc of TEMPO dissolved in solvents of varying polarity, showing the positive linear correlation between EPR hfsc and solvent polarity. Figure 4 shows the variation of EPR hfsc of TEMPO solubilized in aqueous SDS and humic acid micellar solutions above and below the respective cmc. The EPR hfsc decreases above the cmc in SDS solutions as TEMPO partitions into SDS micelles where the polarity is less than water. However, there is no apparent change in the EPR hfsc above the humic acid cmc, estimated to be 0.07% by weight. These results suggest that TEMPO molecules do not penetrate into the humic acid micellar interior but instead adsorb at the surface of humic acid micelles, thereby remaining in continual contact with water. With this understanding of the probable disposition of our hydrophilic and hydrophobic probes, we now consider the results of our paramagnetic relaxation study. The line widths of -9.95 ppm atrazine NMR resonance in 10% humic acid solutions containing either Gd‚EDTA or TEMPO appear in Figure 5. Assuming all of the Gd‚EDTA remains in aqueous solution, Figure 5 represents the Gd‚EDTA concentration as a weighted mole percent of the oxygen atoms in water. Since Gd(III) and TEMPO do not have the same number of unpaired electrons, the concentrations are weighted by their magneton

FIGURE 4. Variation in the electron paramagnetic resonance hyperfine splitting constants for TEMPO (2,2,6,6-tetramethyl-1piperidinyloxy) in aqueous SDS and humic acid solutions. The concentration of SDS or humic acid varies from below to above their respective critical micelle concentrations. The pH of the humic acid solutions were 11.8.

FIGURE 5. Paramagnetic line broadening of atrazine F-19 NMR resonance in 10% aqueous humic acid solution at pH 11.8 as various contents of TEMPO and Gd‚EDTA anion. The mole percent of Gd‚EDTA is computed relative to the oxygen content of water in the solution and weighted by 7.94 Bohr magnetons for Gd(III). The mole percent of TEMPO is computed relative to the combined humic carbon and oxygen content of humic acids in the solution and weighted by 2.00 Bohr magnetons for TEMPO. numbers, 7.94 and 2.00, respectively (57). Assuming all of the TEMPO remains in humic acid micelles, Figure 5 represents the TEMPO concentration as weighted mole percent of the combined carbon and oxygen content, 51.3% and 35.5% (58), respectively, of the humic acids in the solution. Atrazine solubilized in 10% humic acid solutions containing up to 0.04 mol % (i.e., 2.7 mM) Gd‚EDTA exhibits negligible paramagnetic relaxation. TEMPO induces significant paramagnetic relaxation of the -9.95 ppm atrazine resonance, even at TEMPO concentration is as low as 0.0014 mol % (i.e., 0.01 mM).

Discussion Neither solubility enhancement (11, 24, 25) nor molecular aggregation (12-14) prove that concentrated humic acid solutions contain micellar aggregates with hydrophobic interiors. One could imagine solubility enhancement without partitioning if hydrophobic molecules bind to the surface of humic substances. The fractal geometry of humic substances suggest they possess a convoluted structure (14) that may harbor hydrophobic sites at the surface of humic molecules or aggregates of humic molecules. Alternatively, metal ions may form intra- and intermolecular links that knit humic

molecules together creating hydrophobic sites on their surfaces (29, 59). We believe that the differential paramagnetic relaxation of the atrazine F-19 NMR resonance in response to hydrophilic and hydrophobic paramagnetic probes (Figure 5) demonstrates conclusively that hydrophobic domains exist at the interior of humic acid micelles in concentrated humic acid solutions. Neutral organic compounds, such as atrazine, can partition into the hydrophobic humic micellar interior, thereby avoiding paramagnetic relaxation by species confined to aqueous solution. This partitioning presumably accounts for the enhanced apparent solubility. To support our interpretation of these paramagnetic relaxation results, we again refer to the study of glucose binding to a cleft at the surface of an exzyme (40). The paramagnetic broadening of a 5-fluorotryptophan amino acid located in the cleft increases from 7 Hz when the cleft is closed to 130 Hz when the cleft is opened (40). Luck and Falke (40) used Gd‚EDTA concentrations equivalent to those we employed in our study. If atrazine binds to a hydrophobic pocket at the surface of humic micelles, we would anticipate paramagnetic broadening similar to that reported by Luck and Falke (40). The insensitivity of atrazine to paramagnetic relaxation by Gd‚EDTA clearly demonstrates that atrazine solubilized by humic micelles resides at the interior of humic micelles, far removed from the influence of Gd‚EDTA in aqueous solution. The model we propose for atrazine binding to humic substances is analogous to that presented by Nowick and co-workers (60-62). Nucleotide base pairs, formed through cooperative hydrogen bond complexes between weak bases, are not stable in aqueous solution. The hydrophobic domain at the interior of SDS micelles provide an environment suitable for base pairing. The nucleotide base pairs found to exist in SDS micelles (60-62) closely resemble the cooperative hydrogen bond complexes atrazine forms in nonpolar organic solvents (53-56). We believe that atrazine binds to humic substance via cooperative hydrogen bonds that are stable only in the hydrophobic environment found at the interior of humic micelles. Some studies (17, 22) find that fulvic acids exhibit cmc in the same concentration range as humic acids. We cannot conclude from our results that fulvic acid micellar solutions display solubilization and paramagnetic relaxation behavior similar to the humic acids we studied.

Acknowledgments The authors gratefully acknowledge the assistance from NMRFAM staff, Ciba Corporation for providing both radioand fluorine-labeled atrazine. This research was fund by the National Research Initiative-Competitive Grants Program of the USDA (Project 94-37107-0357). Author-Supplied Registry Numbers: Atrazine, 1912-24-9; TEMPO, 2564-83-2; DMSO, 67-68-5; EDTA, 60-00-4; GdCl3, 13450-84-5; (trifluoromethyl)benzene, 98-08-8; D2O, 778920-0.

Literature Cited (1) Lambert, S. M.; Porter, P. E.; Schieferstein, R. H. Weeds 1965, 13, 185-190. (2) Payaperez, A. B.; Cortes, A.; Sala, M. N.; Larsen, B. Chemosphere 1992, 25, 887-898. (3) Lambert, S. M. J. Agric. Food Chem. 1967, 15, 572-576. (4) Lambert, S. M. J. Agric. Food Chem. 1968, 16, 340-343. (5) Chiou, C. T.; Peters, L. J.; Freed, V. H. Science 1979, 206, 831832. (6) Karickhoff, S. W.; Brown, D. S.; Scott, T. A. Water Res. 1979, 13, 241-248. (7) Karickhoff, S. W. Chemosphere 1981, 10, 833-846. (8) McCarthy, J. F.; Roberson, L. E.; Burrus, L. W. Chemosphere 1989, 19, 1911-1920. (9) Chiou, C. T.; Brinton, T. I.; Kile, D. E.; Malcolm, R. L. Environ. Sci. Technol. 1986, 20, 502-508.

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(10) Wershaw, R. L. J. Contam. Hydrol. 1986, 1, 29-45. (11) Wershaw, R. L.; Burcar, P. J.; Goldberg, M. C. Environ. Sci. Technol. 1969, 3, 271-273. (12) Reid, P. M.; Wilkinson, A. E.; Tipping, E.; Jones, M. N. J. Soil Sci. 1991, 42, 259-270. (13) Osterberg, R.; Lindqvist, I.; Mortensen, K. Soil Sci. Soc. Am. J. 1993, 57, 283-285. (14) Rice, J. A.; Lin, J. S. Environ. Sci. Technol. 1993, 27, 413-414. (15) Tschapek, M.; Wasowski, C. Geochim. Cosmochim. Acta 1976, 40, 1343-1345. (16) Tschapek, M.; Scoppa, C. O.; Wasowski, C. Z. Pflanzenernaehr. Bodenk. 1978, 141, 203-207. (17) Chen, Y.; Schnitzer, M. Soil Sci. 1978, 125, 7-15. (18) Rochus, W.; Sipos, S. Agrochimica 1978, 22, 446-454. (19) Tschapek, M.; Wasowski, C.; Scoppa, C. O.; Torres Sanchez, R. M. Agrochimica 1980, 24, 30-38. (20) Tschapek, M.; Wasowski, C.; Torres Sanchez, R. M. Plant Soil 1981, 63, 261-272. (21) Hayano, S.; Shinozuka, N.; Hyakutake, M. J. Jpn. Oil Chem. Soc. 1982, 31, 357-362. (22) Hayase, K.; Tsubota, H. Geochim. Cosmochim. Acta 1983, 47, 947-952. (23) Yonebayashi, K.; Hattori, T. Sci. Total Environ. 1987, 62, 55-64. (24) Shinozuka, N.; Lee, C. Mar. Chem. 1991, 33, 229-241. (25) Guetzloff, T. F.; Rice, J. A. Sci. Total Environ. 1994, 152, 31-35. (26) Kile, D. E.; Chiou, C. T. Environ. Sci. Technol. 1989, 23, 832-838. (27) Kile, D. E.; Chiou, C. T. ACS Symp. Ser.1989, 219, 131-157. (28) Puchalski, M. M.; Morra, M. J.; von Wandruszka, R. Environ. Sci. Technol. 1992, 26, 1787-1792. (29) Engebretson, R. R.; von Wandruszka, R. Environ. Sci. Technol. 1994, 28, 1934-1941. (30) Abragam, A. The Principles of Nuclear Magnetism; Clarendon Press: Oxford, UK, 1962; Chapter 8. (31) Dwek, R. A.; Williams, R. J. P.; Xavier, A. V. In Metal Ions in Biological Systems; Sigel, H., Ed.; M. Dekker: New York, 1974; Vol. 4, pp 61-210. (32) Mazumdar, S. J. Phys. Chem. 1990, 94, 5947-5953. (33) Gao, Z. S.; Kwak, J. C. T.; Wasylishen, R. E. J. Phys. Chem. 1989, 93, 2190-2192. (34) Gao, Z.; Kwak, J. C. T.; Labonte, R.; Marangoni, D. G.; Wasylishen, R. E. Colloids Surf. 1990, 45, 269-281. (35) Wasylishen, R. E.; Kwak, J. C. T.; Gao, Z. S.; Verpoorte, E.; Macdonald, J. B.; Dickson, R. M. Can. J. Chem. 1991, 69, 822833. (36) Yushmanov, V. E.; Imasato, H.; Perussi, J. R.; Tabak, M. J. Magn. Reson. Ser. B 1995, 106, 236-244. (37) Knauer, B. R.; Napier, J. J. J. Am. Chem. Soc. 1976, 98, 43954400. (38) Baglioni, P.; Ottaviani, M. F.; Martini, G. J. Phys. Chem. 1986, 90, 5878-5882. (39) Chung, J. J.; Kang, J. B.; Lee, K. H.; Seo, B. I. Bull. Kor. Chem. Soc. 1994, 15, 198-204.

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(40) Luck, L. A.; Falke, J. J. Biochemistry 1991, 30, 6484-6490. (41) Thurman, E. M.; Malcolm, R. L. Environ. Sci. Technol. 1981, 15, 463-466. (42) Newman, R. H.; Tate, K. R.; Barron, P. F.; Wilson, M. A. J. Soil Sci. 1980, 31, 623-631. (43) McGlamery, M. D.; Slife, F. W. Weeds 1965, 13, 237-239. (44) Gamble, D. S.; Haniff, M. I.; Zienius, R. H. Anal. Chem. 1986, 58, 727-731. (45) Gamble, D. S.; Haniff, M. I.; Zienius, R. H. Anal. Chem. 1986, 58, 732-734. (46) Kalouskova, N. J. Environ. Sci. Health, Part B 1987, B22, 113123. (47) Gamble, D. S.; Khan, S. U. Can. J. Chem. 1988, 66, 2605-2617. (48) Saintfort, R.; Visser, S. A. J. Environ. Sci. Health, Part A 1988, A23, 613-624. (49) Kalouskova, N. J. Environ. Sci. Health, Part B 1989, B24, 599617. (50) Wang, Z. D.; Gamble, D. S.; Langford, C. H. Anal. Chim. Acta 1990, 232, 181-188. (51) Wang, Z.; Gamble, D. S.; Langford, C. H. Anal. Chim. Acta 1991, 244, 135-143. (52) Piccolo, A.; Celano, G.; Desimone, C. Sci. Total Environ. 1992, 118, 403-412. (53) Welhouse, G. J.; Bleam, W. F. Environ. Sci. Technol. 1992, 26, 959-964. (54) Welhouse, G. J.; Bleam, W. F. Environ. Sci. Technol. 1993, 27, 494-500. (55) Welhouse, G. J.; Bleam, W. F. Environ. Sci. Technol. 1993, 27, 500-505. (56) Welhouse, G.; Barak, P.; Bleam, W. F. J. Phys. Chem. 1993, 97, 11583-11589. (57) Ashcroft, N. W.; Mermin, N. D. Soild State Physics, 1st ed.; Saunders College: Philadelphia, 1976; p 826. (58) Sposito, G. The Chemistry of Soils, 1st ed.; Oxford University Press: New York, 1989; p 277. (59) Engebretson, R. R.; Amos, T.; von Wandruszka, R. Environ. Sci. Technol. 1996, 30, 990-997. (60) Nowick, J. S.; Chen, J. S. J. Am. Chem. Soc. 1992, 114, 1107-1108. (61) Nowick, J. S.; Chen, J. S.; Noronha, G. J. Am. Chem. Soc. 1993, 115, 7636-7644. (62) Nowick, J. S.; Cao, T.; Noronha, G. J. Am. Chem. Soc. 1994, 116, 3285-3289.

Received for review March 4, 1997. Revised manuscript received July 28, 1997. Accepted August 8, 1997.X ES9701927 X

Abstract published in Advance ACS Abstracts, September 15, 1997.