Environ. Sci. Technol. 1003, 27,2098-2103
Use of High-Resolution 13C NMR To Examine the Enzymatic Covalent Binding of I3C-Labeled 2,6Dichlorophenol to Humic Substances Patrick G. Hatcher,'*+Jacquellne M. Bortlatynskl,t Robert D. Minard,* Jerzy Dec,g and Jean-Marc Bollagg Fuel Science Program, Department of Chemistry, and Soil Biochemistry Laboratory, Department of Agronomy, The Pennsylvania State University, University Park, Pennsylvania 16802
A novel application of a standard NMR technique has been employed to characterize the enzymatic coupling of a pollutant (2,4-dichIorophenol, 2,4-DCP) to humic material. The application involves the use of 2,4-DCPlabeled with 13C at specific molecular sites in combination with broad-band decoupled 13CNMR to determine the nature of bonding interactions with Minnesota peat humic acid. Any alteration in the chemical structure of the 2,4-DCP due to coupling with the humic acid results in changes in the 13C chemical shifts of the labeled carbons which are indicative of the site and type of bonding interaction. These data provide direct evidence for the formation of covalent bonds between the pollutant and a humic acid via enzymatic coupling. The successof this approach indicates that we now have a method for examining the binding of pollutants and soil organic matter. Introduction Covalent binding of chlorinated aromatic compounds to soil humic material via enzyme-catalyzedredox reactions has been suggested as an important means for mitigating the toxic effects of environmental pollutants (1-4). The presence or addition of oxidoreductive enzymes such as peroxidases or laccases in soil causes free-radical coupling of pollutants to humic material in soil. Studies carried out to examine the enzymatic polymerization of phenolic pollutants such as 2,4-dichlorophenol(2,4-DCP) with itself and with model substrates, which are considered to be analogs of soil organic matter, have shown that covalent bonds are formed in these reactions (4-7). Less direct evidence for enzymatic coupling comes from radiolabeling studies. In these experiments, the inability to extract 14Clabeled 2,4-DCP that has been enzymatically coupled to natural humic substances was interpreted as evidence for covalent bonding (3, 8, 9). The objectiveof this study was to provide direct evidence for enzymatically-induced covalent bonding of a pollutant (2,4-DCP) to humic substances. This evidence was obtained by using a novel application of an NMR method. The method involves the synthesis of 2,4-DCP labeled with 100% 13C at C-2 and C-6 (hereafter referred to as W-2,4-DCP). The 13C-2,4-DCPis then reacted with humic material through an enzyme-catalyzed reaction and examined by 13C NMR. The signals obtained from a l3C NMR experiment are representative of the chemical environment in which the carbon atoms exist, and 13C labeling provides a means of examining selected carbons due to their significantly increased signal intensity. Additional signals at chemical shifts that are different from those of the labeled carbons are indicative of specific + Fuel
Science Program. Department of Chemistry. 8 Soil Biochemistry Laboratory, Department of Agronomy.
t
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Envlron. Scl. Technol., Vol. 27, No.
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types of covalent bonding interactions. Thus, when 13C NMR is used to examine the bonding of pollutants to humic material, not only does the technique provide direct evidence for covalent bonding, but it also provides an indication of the type of bond formed (Le., C-0 vs C-C) between pollutants and humic materials. Experimental Section Materials. Humic acid was prepared by alkali extraction from a Minnesota peat as described by Hatcher et al. (10). The rationale for utilizing this peat humic acid was that a great deal of chemical and spectroscopic information already exists which indicates that the sample appears to be rich in phenolic and carboxylic acid functional groups, which are expected to enhance reactivity with 2,4-DCP. The labeled I3C-2,4-DCP was prepared from phenol labeled with 100% l3C: at the C-2 and C-6 positions (Cambridge Isotope Laboratories, Woburn, MA). A modified version of the chlorination procedure by Ebel et al. (11)was utilized in which the chlorine gas was delivered to the reaction vessel directly rather than the suggested method of chlorine generation from MnOz and HCl. This modification allowed for a more controlled rate of addition of the chlorine gas. The reaction produces a mixture of mono-, di-, and trichlorinated phenols, but by careful selection of reaction conditions, the product yield of 2,4DCP could be maximized to 60%. Complete separation of all the various mono-, di-, and trichlorinated phenol isomers was attained by means of high-performance liquid chromatography on a 4.6- X 150-mm LC-18 (Supelco, Bellefonte, PA). The mobile phase used was 50:50 water/ methanol, adjusted to pH 4 with formic acid. Horseradish peroxidase with an RZ (Reinheitszahl) of 1.2 and an activity of 95 units/mg of solid was purchased from Sigma Chemical Co. (St. Louis, MO). Peroxidase activity is expressed in units defined as the amount of enzyme that forms 1.0 mg of purpurogallin (2,3,4,6tetrahydroxy-5H-benzocyclohepten-5-one) from pyrogallol in 20 s at pH 6 and 20 "C. Binding of 13C-2,4-DCPto Minnesota Peat Humic Acid. Since the Minnesota peat humic acid is only partly soluble in water at pH 7.2, a saturated solution was prepared before addition to the 13C-2,4-DCPmixture. To accomplish solubilization, the humic acid was first suspended in pH 7.2 universal buffer (0.2 M acetic acid, 0.2 M boric acid, 0.2 M phosphoric acid, and 1 N NaOH) at a concentration of 2 mg/mL. The pH of the solution was raised to greater than pH 10 with 40% NaOH, and the mixture was placed in a sonicator for 30 min. The volume of the solution was tripled with a universal buffer, and the solution was readjusted to pH 7.2. The resulting suspension was centrifuged, and the supernate was diluted 1.7 times with the buffer. Aliquots of this solution were then used for the following binding procedure. A 8.4-mL volume of 13C-2,4-DCP solution in pH 7.2 universal buffer (0.4 mg/mL) was placed in a 250-mL 0013-936X/93/0927-2098$04.00/0
0 1993 American Chemical Society
Erlenmeyer flask and diluted with 75 mL of the buffer along with 14C-labeled 2,4-DCP (0.67 pCi in 70 pL of methanol) and 1.9 mg of humic acid in 11.7 mL of the buffer. The ratio of humic acid to 2,4-DCPwas determined on the basis of the maximum solubility of the humic acid and the minimum amount of 2,4-DCP that was needed to attain a reasonable signal-to-noise ratio. The mixture was then treated with 64 units of peroxidase (236 pL of water containing 2.9 mg of the enzyme per 1mL) and 139 pL of 1%H202 and incubated for 2 h. After incubation, the reaction mixture was acidified with HCl to pH 1, stored overnight in a cold room for complete precipitation of humic acid, and centrifuged. The pellet was washed several times with acidified water (pH 1) and was then dissolved in 1 mL of 1%NaOD and subjected to NMR analysis. Analysis of Radioactivity. 14C-Labeled2,4-DCP was bound to Minnesota peat humic acid by the method described above. The pellets of the precipitated humic acid with the attached 14C-labeled2,4-DCP were washed three times with acidified water, dried, washed twice with dichloromethane, dried, and washed twice with methanol to remove loosely bound 13C-2,4-DCPand dissolved in NaOH. Aliquots from the washings and the dissolved pellet were then placed in Scinti-Verse I1 liquid scintillation cocktail (Fisher Scientific Co., Fair Lawn, NJ) for 14Ccounting. The radioactivity was measured with a Beta Trac 6895 liquid scintillation counter (Tracor Analytic, Elk Grove Village, IL). Aliquots of the extracted solutions were analyzed by radiocounting, which revealed that 53.9 % of the initial radioactivity became bound to humic acid during the enzymatic reaction. 13CNuclear Magnetic Resonance Techniques. The 13C NMR inverse-gated spectrum of 13C-2,4-DCP was obtained using composite pulse decoupling on a Bruker AM500 NMR spectrometer with a resonance frequency for 13Cof 125 MHz. A pulse angle of 45O and a relaxation delay of 2 s were used with a sweep width of 31 000 Hz which results in a 250-ppm chemical shift range. Free 13C-2,4-DCPwill have long spin-lattice relaxation times of about 2 s and may exhibit signal saturation. Covalently bound 13C-2,4-DCP should exhibit relaxation behavior similar to that of humic acid itself [i.e., short spin-lattice relaxation times of less than 2 s, Preston and Blackwell (1213;therefore, the use of minimal relaxation times between pulses should not saturate the signals or distort the signal intensities for bound 13C-2,4-DCP. Approximately 23 000 acquisitions were required for suitable spectra. The sample of 13C-2,4-DCPused for chemical shift measurements was prepared in a universal buffer containing 10% DzO at neutral pH. The same sample was used to determine chemical shifts at pH 10by dropwise addition of 40 % aqueous NaOD to achieve the proper pH.
Results and Discussion Labeling Studies. Although I4C radiolabeling cannot be used to examine structural changes, it provides quantitative information on the mass balance of the reaction and on how the 13C-2,4-DCP is partitioned between extractable and nonextractable components. In the presence of horseradish peroxidase, 54 % of the initial radioactivity remained with the solid pellet isolated from the first precipitation of the humic acid. Table I contains the data showing recoveries of 14C from subsequent
Table I. Percent Distribution of Radioactivity following the Extraction of I4C-Labeled2,4-DCP Bound to Minnesota Peat Humic AcidP sample
radioactivity (dpm)
radioactivity (%)
extraction with acidified water extraction with dichloromethane extraction with methanol humic acid after extractions total recovery
19 510 16 385 30 305 119 573 185 773
10.3 8.6 16.0 63.1 98.0
a The initial radioactivity incorporated in the humic acid amounted to 189 500 dpm.
a
120
I
ppm
130
,
I 200
I ppm
I
1
I 100
I
I
I 25
Figure 1. 13CNMR spectrum of l3C-2,4-DCPadded to Minnesota peat humic acid at basic pH in the absence of horseradish peroxidase. (a) Expanded region 110-140 ppm. (b) Full spectrum 0-200 ppm.
extractions of the pellet with acidified water, dichloromethane, and methanol as well as the final content of 14C activity in the isolated humic acid. Approximately 63% of the initial radioactivity attached to the humic acid by the horseradish peroxidase remained after repeated extractions. This is the percentage of 14C-labeled 2,4DCP that is thought to be covalently bound to the humic acid, since the water extraction should haveremoved any loosely bound organicmolecules while the dichloromethane and methanol extractions should have removed any low molecular weight hydrophobic and/or polar organic molecules. The acidified water extractions removed 10% of the loosely bound 14C label, while the dichloromethane and methanol extractions removed an additional 8.6% and 16%, respectively. The solvent extractions may also remove some of the humic acid, so all of the radioactivity found in these solutions may not be due entirely to free 2,4-DCP. In order to avoid sample losses in experiments where 13C-2,4-DCPwas used, the organic solvent extractions were eliminated, and the humic acid was extracted repeatedly only with acidified water. l3C NMR Studies. The 13C NMR spectrum of 13C2,4-DCP in the presence of Minnesota peat humic acid under basic conditions is shown in Figure 1. The spectrum was obtained at basic pH, because this humic acid is only Environ. Scl. Technol., Vol. 27, No. 10, 1993 2099
Table 11. 18CChemical Shifts of 2,g-Labeled 2,4-DCP in the Presence and in the Absence of Humic Acid
pH 11 7 11
I
humic acid present
chemical shift of the C-6 carbon (pprn)
chemical shift of the C-2 carbon (ppm)
no no Yes Yes
119.95 117.13 119.91 117.13
123.10 120.48 123.10 120.41
sparingly soluble at pH 7. Under basic conditions, the phenol exists as the phenolate anion. The anion distributes its negative charge throughout the aromatic ring, and the chemical shifts of the C-2 and C-6 labeled carbons increase 2-3 ppm due to the associated increase in electron density (13). Table I1 lists the 13Cchemical shifts of the C-6 and C-2 labeled carbons for 13C-2,4-DCPat neutral and basic PH. On the basis of 14C labeling experiments, little or no covalent binding is expected in the absence of oxidoreductive enzymes. Similar results were observed in a 13CNMR experiment in which the Minnesota peat humic acid in the absence of the oxidoreductive enzyme is allowed to interact with 13C-2,4-DCP. The resulting l3C NMR spectrum (Figure 1) contained only two signals at 119.95 and 123.10ppm, which were assigned to the C-6 and C-2 carbons of 13C-2,4-DCPphenolate ion, respectively. These signals are two sets of doublets (JCCapproximately 15 Hz) which result from the two-bond coupling of the labeled carbons. Another point of significanceis that the spectrum is devoid of any signals resulting from the unenriched (1% natural abundance) carbons of the W-2,4-DCP or the humic acid whose signals are too weak to be observed in the number of scans used in these experiments. The 13CNMR spectrum of 13C-2,4-DCPbound to humic acid is shown in Figure 2. Figure 2c illustrates the entire spectrum from 0 to 200 ppm, while Figures 2b and 2a illustrate expanded portions of the spectrum from 100 to 140 ppm and 140 to 200 ppm, respectively. In striking contrast to the spectrum of free 13C-2,4-DCP(Figure 11, the spectrum of W-2,4-DCP incubated with humic acid and horseradish peroxidase (Figure 2) displays a large dispersion of 13C chemical shifts caused by the covalent bonding of the 13C-labeledcarbons to the humic acid. The chemical shifts were expected to be distributed throughout the aromatic region of the spectrum, because the humic acid macromolecular structure contains a variety of functionalities capable of forming covalent bonds to 13C2,4-DCP. Each type of covalent bond exhibits a slightly different effect on the electronic environment surrounding the labeled carbons, which are then detected as changes in the 13C chemical shifts of the labeled carbons (13). The only similarity between the spectra in Figures 1 and 2 is the presence of two large doublets at 119.9and 123.1ppm. The chemical shifts of these two doublets correspond to those for free (unbound) 13C-2,4-DCPat basic pH. The presence of unbound '3G-2,4-DCP may result from incomplete extraction by water or from binding by a mechanism other than covalent bonding. Humic acids are known to be hydrophobic in character and trap smaller molecules within their macromolecular structures (14). Lee et al. (15, 16) and Schwarzenbach et al. (17) have shown that chlorinated phenols associate with soil organic matter through a weak noncovalent interaction. In addition, Lee et al. (15) were able to show that the 2100
Envlron. Scl. Technol., Vol. 27, No. 10, 1993
a
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Flgure 2. 13CNMR spectrum of l3C-2,4-DCP enzymatically bound to Minnesota peat humicacid after treatment with horseradishperoxidase. (a) Expanded region 140-180 ppm. (b) Expanded aromatic region 110-140 ppm. (c) Full spectrum 0-200 ppm.
phenolate analogs of pentachlorophenol, 2,4,5-trichlorophenol, and 2,3,4,5-tetrachlorophenoldisplay similar behavior. Thus, the 13C-2,4-DCP might actually be trapped in the humic acid structure and, therefore, not be extractable with acidic water. Another explanation for the presence of free 13C-2,4-DCPis that the NaOD may hydrolyze any weakly bonded 13C-2,4-DCP. The question of whether the dispersion of signals observed in Figure 2 could be due to this free 13C-2,4-DCP associating with the humic acid through dipole, hydrogenbonding, or hydrophobic attractions must also be considered. As discussed in a previous section, the 13CNMR spectrum, shown in Figure 1,clearly illustrates the absence of peaks other than those of the lW-2,4-DCP. If the l3C2,4-DCP is only weakly associating with the humic acid by one of the mechanisms described above, then these types of interactions are apparently unable to induce a dispersion of chemical shifts. Thus, this 13C NMR technique cannot be used to identify weakly associated interactions with the Minnesota peat humic acid. The appearance of two intense sets of apparent triplets at 123.7 (center of region E) and 131.4 ppm (center of region G) in Figure 2 suggests that 13C-2,4-DCPunits may be linked to form an oligomeric moiety (substructure I). The apparent coupling pattern (triplets) can only arise from the close proximity of two sets of '%-labeled carbons and is probably the result of the overlap of two doublets (a doublet of doublets) from the two-bond coupling of the 13C-labeledcarbons at C-6 and C-2' (substructure I). This type of l3C-'3C two-bond coupling is rarely observed in natural abundance l3C NMR spectra because of the low probability of having two adjacent 13Cs.
0
I,#
6' 118.8
lZ2'O
Cl
r The dimeric moiety (substructure I) was assigned to the multiplets at 131and 123ppm in Figure 2 on the basis of calculated l3C chemical shifts. Lacking appropriate model compounds, calculated chemical shifts are often used for the assignment of signals in the NMR spectrum (13). The calculated 13C chemical shifts were obtained using the 13C-2,4-DCP phenolate as the base structure, because (1) the proposed 2,4-DCP dimer moiety would exist in base as a phenolate anion or dianion and (2) the best estimates of chemical shifts will be obtained from the base set needing the least number of additions, i.e., the substructural features of the dimer moiety are most closely related to 2,4-DCP. The first doublet of doublets at approximately 131ppm is tentatively assigned to the twobond coupling of C-6 to both C-2 and C-2', while the other doublet of doublets at 123 ppm is tentatively assigned to the two-bond coupling of C-2' to both C-6 and C-6'. The observation of this 2,4-DCP dimer substructure led us to carry out a control experiment that might allow differentiation of oligomerized 2,4-DCP from humic acidbound 2,4-DCP. Reaction of 13C-2,4-DCP catalyzed by an oxidoreductase in the absence of humic acids did indeed yield a polymerized product showing a dispersion of chemical shifts indicative of a polymer (Hatcher et al., manuscript in preparation). However, this reaction is always accompanied by precipitation of 2,4-DCP oligomers, whereas no precipitation or even increased turbidity was observed for the reaction with humic acids. The resulting 13CNMR spectrum of the polymerized product (obtained in chloroform-d due to lack of solubility in NaOD) has signals in regions that do not correlate with signals in the humic acid-bound 13C-2,4-DCPspectrum, even allowing for a 2-3-ppm variation due to solvent. In addition, this control spectrum lacked multiplet signals at 131and 123 ppm that are found in the humic acid-bound 13C-2,4-DCP spectrum and that are attributed to substructure I. The remaining signals in Figure 2 consist of additional sharp multiplets or singlets which appear at approximately 118,122,127.9,133,167,and 172 ppm. The latter signals at 167 and 172 ppm are assigned to natural abundance carbons in carbonate and bicarbonate ions. In addition to these sharp signals, broad peaks are also observed. In particular, the region between 123 and 125 ppm contains a broad envelope of resonances superimposed on the sharp signals discussed above. Also, broad peaks are observed between 127 and 130 ppm as well as between 136 and 138 ppm. The complexity of the signals reflects the numerous sites that are available for covalent bonding in the humic acid macromolecular structure. Specific assignments for the types of bonding responsible for these complex signals are discussed below. Another source of signals in the NMR spectrum that must be considered are those which may be present as the
result of unlabeled carbons. It is unlikely that any of the signals in Figure 2 arise from the carbons of humic acid or unlabeled carbons of 2,4-DCP, which are present at natural abundance levels (l.l%), for three reasons: (1) the spectrum shown in Figure 1 clearly illustrates that carbons of the 13C-2,4-DCPpresent at natural abundance levels are not observed under the given spectral conditions; (2)the presence of doublet species supports the conclusion that only 13C-2,4-DCP labeled carbons appear in the spectrum (as mentioned above, doublets are rarely observed when l3C is present only in natural abundance); and (3) the natural abundance 13C NMR spectrum of Minnesota peat humic acid contains abundant signals in the aliphatic region, and no signals are observed in this region in Figures 1and 2, indicating that only I3C-labeled carbons are present in the I3C spectrum of unbound or bound W-2,4-DCP. Structural Assignments. The complexity of the NMR signals in Figure 2 and the lack of any model compound data for comparison make the task of assigning all the peaks quite complex. A large variety of carboncarbon and carbon-oxygen bond types can be formed between the 13C-2,4-DCPand a macromolecular structure such as humic acid. One reasonable approach for tentatively assigning the complex resonances in such a spectrum is to estimate the chemical shifts of the C-2 and C-6 carbons when the 13C-2,4-DCPforms bonds to oxygen or the ring carbons to yield various bonding types such as ethers, esters, and alkyl-aryl or aryl-aryl carbon-carbon bonds between 2,4-DCP and humic acid. Figure 3 presents a number of the possible types of covalent bonding interactions that would be expected to occur between 2,4-DCP and humic acid. These interactions were chosen on the basis of the reactions observed by Minard et al. (5)in the laccase-catalyzedoligomerization of 2,4-DCP and the rational chemistry predicted for reactions of phenols with humic acid functional groups: ester formation, ether formation, and oxidative C-C and C-0 coupling, with reactions most likely occurring at the ortho and para positions or the phenolic oxygen. For each of these structures, the 13Cchemical shifts of the C-2 and C-6 carbons were calculated using generally accepted NMR additivity rules and the known chemical shifts of C-2 and C-6 carbons of 13C-2,4-DCPor its phenolate as reference points. For the carbon-carbon linkages (structures 1-3) and the ether linkages (structures 4-7), the calculated shifts are slightly different depending on whether the linkages are to aliphatic (shown in plain text) or aromatic (shown in italic text) carbons in the humic acid. In the case of the ester linkages (structures 8-11), the chemical shift was virtually the same for either alkyl or aryl humic acid substituents and so only one value is shown. In the case of structures 1,2,5,6,9, and 10, the attachment to humic acid involves the loss of chlorine at C-2 or C-4 of the 2,4DCP ring. Dechlorination of chlorophenols has previously been observed during enzymatic polymerization reactions (4, 5).
As one would expect for the carbon-carbon-linked compounds 1-3, the calculated changes in chemical shifts are greatest when the new bond is formed directly at C-2 (1) or C-6 (31, in which case the values shift downfield 10-15 ppm to the 130-135-ppm region. Calculated changes in chemical shifts are even more extensive if the new bond to C-2 or C-6 is to oxygen: for ethers 5 and 7, the chemical shifts are calculated to be in the 146-151-ppm range, and Environ. Sci. Technol., Vol. 27, No. 10. 1993 2101
Table 111. Tentative Assignment of Chemical Shift Regions for I3C-2,4-DCPBound to Minnesota Peat Humic Acid b Chemical shifts for the
phenolate of 2,4-DCP
&I
148.3
chemical shift range (pprn)
A B C D
118-119.5 119.5-121 121-122 122-123.5
E F G H I J
123.5-125 126-129 130-133 133-134 135-138 146-148
labeled 2,4-DCP site C-6 C-6 c-2 c-2 C-6 c-2 c-2 c-2 C-6 C-6 c-2
structure(s) in Figure 4 I,2
4,5,6, 10 2 3,6, 10 8 394
8 1 3 3 5
I
O(humlc)
119.6 120.2
bI
bl
5
7
CARBONOXYGEN BINLXNG VIA AN ETHER LINKAGE
B p'C'C(humlc)
61 9
CI
11
CARBONOXYGEN BINDING VIA AN ESTER LINKAGE
Figure 3. Suggested carbon-carbon and carbon-oxygen bonds of 13C-2,4-DCP bound to Minnesota peat humic acid. The calculated shifts for the C-2 and C-0 labeled carbons of the 2,4-DCP when bound to an aliphatic and aromatic (in italics) carbon of humic acid are listed. When bonds toaromatic or aliphatic humic acid moietles yield chemical shlfts that are identicalor very close, only one chemlcaishift is reported, and this is shown in bold.
for esters 9 and 11,in the 140-143-ppm range. Otherwise, for all the other structures where bonds are formed at C-4 or the phenolic oxygen, the calculated chemical shifts for C-2 and C-6 vary only a few ppm from those observed for unreacted 13C-2,4-DCP(119.9and 123.1ppm, respectively) and therefore would tend to cluster in the 115-127-ppm range. Using these calculated chemical shift values, the NMR spectrum of humic acid-bound W-DCP shown in Figure 2 can now be examined in terms of the potential presence or absence of chemical linkages of the types represented by structures 1-11. Table I11 contains tentative assignments of the selected chemical shift regions, designated A through J, in which the signals correspond to the calculated chemical shifts for many of the predicted chemical linkages shown in Figure 3. The data indicate that all but structures 4 (in which the humic acid substituent is alkyl), 7,and 11 are compatible with the observed NMR data. In the case of structure 4, an ether linkage through the phenolic carbon to an aliphatic carbon of humic acid is probably not occurring, because the l3C chemical shift for C-6 carbon under such conditions (approximately 115ppm) should be found in the region of the 13C spectrum (Figure 2) where no resonances are observed. The same type of ether linkage to an aromatic carbon in the humic acid only 2102
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Envlron. Sci. Technol., Vol. 27, No. 10, 1993
slightly increases the chemical shifts of the C-2 and C-6 carbons. It is difficult to determine whether such structures exist, because the signals would be buried under signals due to unbound 2,4-DCP. Carbon-oxygen bonds formed at the C-6 position via an oxygen-containing functional group of the humic acid (structure 7) shift the signals of the labeled substituted carbons of the 2,4-DCP to the chemical shift range of 143151 ppm. Again, it is unlikely that such linkages exist, because no peaks are observed in Figure 2 in this region. An ether linkage formed at the C-4 position to the humic acid (structure 6) results in only a slight change in the chemical shifts of the C-2 and C-6 labeled carbons. Here, too, we cannot discern whether such linkages exist because of the overlap of peaks with those of unbound 2,4-DCP. Finally, the covalent linkage represented by structures 8-11 in Figure 4 results from carbon-oxygen bonds which form the alkoxy groups of esters. When the ester linkage is at the C-2 or C-6 carbons, the chemical shifts of the substituted carbons increase and appear in the 139-143ppm range. Again no peaks are observed in this region of Figure 2, suggesting the lack of these structures. An ester linkage through the C-1 carbon of the 13C-2,4-DCP (structure 8) increases both the C-2 and C-6 carbon chemical shifts. These chemical shifts are in regions of the spectrum (regions C, D, and F in Figure 2) where significant intensity is observed, suggesting that a large proportion of the 2,4-DCP is bound to humic acids by such a linkage. Conclusions Until now, the only evidence for the covalent bonding of 2,4-DCP to humic acid relied upon 14C labeling and the structural information gained from model compound studies (2-9). 13Clabeling in conjunction with 13CNMR has provided a direct method for the examination of the bonding of 13C-2,4-DCP to a complex macromolecular structure such as humic acid. The structural changes were detected directly by observing the concomitant changes in chemical shifts of the 13C-2,4-DCPcarbons as the humic acid covalently binds to the 2,4-DCP. In the absence of enzymes, we detected only the labeled carbons of the free, unbound 13C-2,4-DCPboth in the presence and in the absence of humic acid. No signals for the humic acid carbons are observed because they exist at only natural abundance levels. When the reaction of 13C-2,4-DCPwith humic acid is performed in the presence of oxidoreductive enzymes, a large number of peaks are observed in the NMR
spectrum, reflecting the dispersion of chemical shifts induced by covalent bonding of 13C-2,4-DCPto itself and to humic acid. These new l3C chemical shifts have been tentatively assigned by comparison to calculated chemical shifts for the various types of carbon-carbon and carbon-oxygen linkages that can be reasonably postulated. Chemical shift additivity rules have been applied to structures which are predicted on the basis of the prior studies of Minard et al. (5) and the known reactivities of phenols in general. Thus, the various peaks observed in the spectrum of enzymatically bound W-2,4-DCP are suggested to represent (1)2,4-DCP polymerized to itself, (2) 2,4-DCPbound to humic acid through ester linkages, (3) 2,4-DCP bound to humic acid through phenolic ether linkages, and (4) 2,4-DCP bound to humic acid through carbon-carbon linkages. Many of the observed peaks can only be explained by reaction at chlorinated sites of the 2,4-DCP, indicating that removal of the chlorine atom must have preceded or been coincident with the binding to humic acid. While many possible bonding interactions can be ruled out because concordant peaks are not observed in the spectrum of 2,4-DCP bound to humic acid, we do note that two bonding interactions yield the strongest peaks. Bonding through an ester linkage at C-1 (structure 8) and carbon-carbon bonds at C-6 (structure 3) and C-4 (structure 2) appear to be the most likely types of interactions. A more precise assignment of the chemical shifts will require further synthetic work in which the reference 13C chemical shifts are derived from the spectra of appropriate model compounds. It is clear that the NMR approach taken in this study has great potential for the investigation of the binding of pollutants to humic materials. Although these studies involve 2,4-DCP and a specific humic acid, a similar methodology can be employed with other pollutants and humic materials to provide much information on the binding of pollutants to soil organic matter.
Acknowledgments We would like to thank William R. Wilkinson and Faezeh Firouzkouhi for the preparation of the 13C-labeled 2,4-
DCP. We would also like to acknowledge the College of Agriculture Intercollege Research Grants Program for their support.
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Received for review November 30, 1992. Revised manuscript received May 31, 1993. Accepted June 16, 1993.' ~~
Abstract published in Advance ACS Abstracts,August 15,1993.
Environ. Sci. Technol., Vol. 27, No. I O , 1993 2103
