Environ. Sci. Technol. 2000, 34, 4278-4286
Sources and Haloacetic Acid/ Trihalomethane Formation Potentials of Aquatic Humic Substances in the Wakarusa River and Clinton Lake near Lawrence, Kansas M I C H A E L L . P O M E S , * ,† CYNTHIA K. LARIVE,‡ E. MICHAEL THURMAN,† W. REED GREEN,§ WILLIAM H. OREM,| COLLEEN E. ROSTAD,# TYLER B. COPLEN,| BENJAMIN J. CUTAK,‡ AND ANN M. DIXON‡ U.S. Geological Survey, 4821 Quail Crest Place, Lawrence, Kansas 66049, Department of Chemistry, The University of Kansas, Lawrence, Kansas 66045, U.S. Geological Survey, 401 Hardin Road, Little Rock, Arkansas 72211, U.S. Geological Survey, 956 National Center, 12201 Sunrise Valley Drive, Reston, Virginia 22192, and U.S. Geological Survey, 408 Federal Center, Denver, Colorado 80225-0046
Gram quantities of aquatic humic substances (AHS) were extracted from the Wakarusa River-Clinton Lake Reservoir system, near Lawrence, KS, to support nuclear magnetic resonance (NMR) experimental studies, report concentrations of dissolved organic carbon (DOC) and AHS, define sources of the AHS, and determine if the AHS yield sufficient quantities of haloacetic acids (HAA5) and trihalomethanes (THM4) that exceed U.S. Environmental Protection Agency (EPA) Maximum Contaminant Levels (MCL) in drinking water. AHS from the Wakarusa River and Clinton Lake originated from riparian forest vegetation, reflected respective effects of soil organic matter and aquatic algal/bacterial sources, and bore evidence of biological degradation and photodegradation. AHS from the Wakarusa River showed the effect of terrestrial sources, whereas Clinton Lake humic acid also reflected aquatic algal/bacterial sources. Greater amounts of carbon attributable to tanninderived chemical structures may correspond with higher HAA5 and THM4 yields for Clinton Lake fulvic acid. Prior to appreciable leaf-fall from deciduous trees, the combined (humic and fulvic acid) THM4 formation potentials for the Wakarusa River approached the proposed EPA THM4 Stage I MCL of 80 µg/L, and the combined THM4 formation potential for Clinton Lake slightly exceeded the proposed THM4 Stage II MCL of 40 µg/L. Finally, AHS from Clinton Lake could account for most (>70%) of the THM4 concentrations in finished water from the Clinton Lake Water Treatment Plant based on September 23, 1996, THM4 results.
Introduction Aquatic humic substances (AHS) have been subjected to various nuclear magnetic resonance (NMR) techniques (1H, 13C, 31P, and 15N) to gain information on the types and numbers of constituent functional groups (1). As defined by 4278
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Thurman (2), AHS are colored, polyelectrolytic, nonvolatile organic acids that range in molecular weight from 500 to >10,000 daltons and are isolated from water by sorption onto XAD resins or comparable procedures. Humic acids consist of components of AHS that precipitate when the sample pH is decreased to 1.0, whereas fulvic acids remain in solution. With the protonation of most carboxylic acid groups at pH 1.0, the more hydrophobic humic acids precipitate because of their smaller carboxylic acid content and greater molecular weight. Overall, AHS commonly constitute about 50% of the dissolved organic carbon (DOC) in a water sample filtered through a 0.45-µm filter. Recently, researchers have developed novel techniques involving NMR to examine the physical and structural properties of AHS on finer scales. The combination of spectral editing techniques and modified pulsed-field gradient experiments involving proton NMR-derived diffusion coefficients (3) of the Suwannee River (GA) fulvic acid standard (4, 5) yielded radii of gyration comparable to values reported by small-angle X-ray scattering (6, 7). Additionally, the use of 113Cd NMR has allowed the measurement of Cd2+ exchange rates for Suwannee River fulvic acid (8, 9). Experiments with these NMR techniques, as well as solidstate 13C NMR, on previously uncharacterized AHS, like those in the Wakarusa River-Clinton Lake Reservoir system near Lawrence, KS, have the potential to consume relatively large quantities of sample mass (75-200 mg per experiment). Additionally, at least 500 mg per sample can be consumed by classical characterization techniques that include elemental analysis, titration, Fourier transform infrared spectroscopy (FTIR), δ13C determination, and determination of lignin oxidation products (LOP) (2). Such characterization techniques yield detailed information about the source of AHS in lake and reservoir settings (10-12), and whether water percolated through the soil (13) prior to reaching the river or reservoir. One newer characterization technique, the determination of haloacetic acid (defined to include mono-, di-, trichloracetic, mono-, and dibromoacetic acid; HAA5) and trihalomethane (defined to include bromodichloromethane, dibromomonochloromethane, bromoform, and chloroform; THM4) formation potentials of AHS (11, 12) can determine if AHS yield sufficient concentrations of HAA5 and THM4 to raise regulatory concerns under the Disinfection Byproducts Rule (14). Thus, gram quantities of AHS are required to support NMR experiments and characterization efforts. This paper describes the extraction of gram quantities of AHS from the Wakarusa River-Clinton Lake Reservoir system, near Lawrence, KS, reports concentrations of DOC and the proportions of DOC accountable as AHS, defines sources of the AHS, and presents a determination of whether the HAA5 and THM4 formation potentials of the AHS are sufficient to create regulatory concerns. The September 10October 10 time of sample collection took place before any appreciable leaf-fall occurred from deciduous trees and represents a time of minimal terrestrial input to the riverreservoir system. Work in other settings has shown that source * Corresponding author phone: (785)296-6372; fax: (785)832-3500 (U.S. Geological Survey); e-mail:
[email protected]. Present address: Kansas Department of Health and Environment, Bureau of Environmental Remediation, Forbes Field, BLDG 740, Topeka, KS 66620-0001. † U.S. Geological Survey, Lawrence, KS. ‡ The University of Kansas. § U.S. Geological Survey, Little Rock, AR. | U.S. Geological Survey, Reston, VA. # U.S. Geological Survey, Denver, CO. 10.1021/es991376j CCC: $19.00
2000 American Chemical Society Published on Web 09/07/2000
FIGURE 1. Location of sampling sites on Wakarusa River and in Clinton Lake near Lawrence, KS. material and time of year can affect amounts of natural organic matter as well as its reactivity and ability to generate disinfection byproducts (DBP) (15). Thus, AHS collected during this time could constitute baseline against which the effects of other hydrologic and organic inputs can be evaluated. Completed in 1980, Clinton Lake occupies 28.4 km2 (multipurpose pool of 233 m above sea level) (16) of the 1100-km2 Wakarusa River watershed (17) (Figure 1). The Wakarusa River watershed is located in the Osage Section of the Central Lowland physiographic province which consists of gently rolling, westwardly dipping plains of low relief interrupted by east facing escarpments (18). Near Lawrence, these plains are broken by broad valleys occupied by the Wakarusa and Kansas Rivers. Land use in the watershed consists mainly of cropland with some grazing land (19), and riparian gallery forests line the Wakarusa River. A U.S. Geological Survey (USGS) gaging station (no. 6891478) is located in the spillway house of Clinton Dam. Data collected at the station include hourly precipitation measurements and values of lake elevation relative to sea level. No other stream gages are located on the Wakarusa River upstream of Clinton Lake.
Experimental Procedures Sample Collection. Between September 10 and October 10, 1996, 540 L of water were collected from the Wakarusa River, and 610 L of water were collected from Clinton Lake (Figure 1) during 12 sample-collection events. Sample collection began with the deployment of a 5.1-cm diameter polyvinylchloride well screen (0.25-mm slots) and support framework in less than 1 m of water. Polyethylene tubing (6.4-mm diameter) connected the well screen to the peristaltic pump drive (Masterflex 07521 Series, Cole-Parmer, Chicago, IL)
fitted with C-Flex tubing (size 15, Cole-Parmer, Chicago, IL) and powered by a portable generator. Water was continuously pumped into a beaker until pH (Model SA 250, Orion Research, Cambridge, MA) and specific conductance (Model 604, Amber Scientific, Eugene, OR) values stabilized. At stabilization, a 0.45-µm AquaPrep 600 capsule filter (Gelman Sciences, Ann Arbor, MI) was connected to the pump outlet and rinsed with approximately 1 L of water. In tests with distilled water, rinsing the filters with 1 L of water reduced the DOC concentrations to the extent that they were indistinguishable from organic carbon concentrations attributable to distilled water blanks. After rinsing the filter, water was pumped into a 20-L glass carboy. The process was repeated until three carboys (about 60 L total) were collected from each sampling site and transported to the USGS laboratory in Lawrence, KS, where they were stored in a walkin refrigerator (4 °C) prior to extraction of AHS that evening. When suspended-sediment loads in the Wakarusa River made onsite filtration impractical, water was transported to the USGS laboratory in 20-L polypropylene carboys (Nalge Company, Rochester, NY). There water was filtered with glassfiber filters (rinsed with 1 L of river water) through a 142-mm diameter, stainless steel plate filtration unit (Millipore, Bedford, MA) connected to a 20-L stainless steel tank. Pressurizing the tank with N2 forced the sample through the filter unit. The filtrate was collected in a second polypropylene carboy from which a peristaltic pump drew water for filtration with a 0.45-µm capsule filter. Water filtered with the 0.45-µm capsule filter was collected in 20-L glass carboys prior to acidification and extraction of AHS. During each sample-collection event, water from sampling sites 1 and 2 was collected in 125-mL, amber glass bottles for DOC determinations using an organic carbon analyzer (Model TOC-700, Oceanographic Instruments, College StaVOL. 34, NO. 20, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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tion, TX). Water in the 125-mL bottles remaining after DOC analyses (approximately 95 mL) was saved to make composite samples from each site. Subsequent organic carbon determination of the sample composites then gave DOC concentrations representative of the 540 L of water collected from the Wakarusa River and 610 L collected from Clinton Lake. Extraction and Fractionation of AHS. Following each sample-collection event, water from the three chilled 20-L carboys from both sites was acidified to pH 2.0 with concentrated HCl and pumped into two 1.2-L columns (one for the river water and one for the lake water) filled with XAD-8 resin (Rohm & Haas, Philadelphia, PA) following the procedure of Thurman and Malcolm (20). Adsorbed AHS were recovered from XAD-8 columns by back elution with 0.1 N NaOH and collected in 2-L glass bottles. The pH of the NaOH eluate was lowered to 2.0, and a 40-mL aliquot of the desorbed and acidified AHS was collected for DOC analysis. Additional pH adjustment to 1.0 and chilling the bottles facilitated the fractionation of AHS to humic and fulvic acids. Precipitated humic acids were separated by vacuum filtration through pre-rinsed 47-mm diameter glass-fiber filters (P/N 66078, Gelman Sciences, Ann Arbor, MI) using a glass filter holder and sidearm flask. Spent filters containing accumulated humic acids were set aside, and the fulvic-acid filtrate was applied to the larger of two reconcentration columns filled with XAD-8 resin. Prior to application of the fulvic-acid filtrate to the column, a second 40-mL aliquot was collected for DOC analysis. Filters containing humic acid were placed on the vacuum apparatus. Several 100-mL rinses of 0.1 N NaOH were drawn through the filter (under vacuum) until the humic acids were removed. The humic-acid filtrate was acidified and applied to the smaller of two reconcentration columns filled with XAD-8 resin. Procedures including the elution, cation exchange, and lyophilization of AHS have been described elsewhere (12, 13). After each freeze-drying run, lyophilized humic and fulvic acid samples were stored in labeled glass vials. In effect, these humic and fulvic acids constituted composite samples representing 540 L of water collected from the Wakarusa River and 610 L collected from the Wakarusa River. Characterization of AHS. Several analytical techniques were used to characterize the AHS composite samples collected from the two sites: 13C NMR, δ13C determination, titrations, determination of lignin oxidation products (LOP), elemental analysis, Fourier transform infrared spectroscopy (FTIR), and determinations of HAA5 and THM4 formation potentials. Depending on the amount of sample available, 75 to 200 mg of the lyophilized humic and fulvic acids were weighed into individual centrifuge tubes in preparation for 13C NMR. The humic- and fulvic-acid samples were dissolved in 3.0 mL of 0.1-M NaOD (Isotec, Miamisburg, OH) in D2O (Cambridge Isotope Laboratories, Andover, MA) with additional base added as necessary to solubilize the sample. Because D2O solutions were used in this work, the acidity of these solutions was reported as pD to correct for the isotope effect at the glass electrode (21). If the pD of humic and fulvic acid solutions was high following the solubilization procedure, DCl (Sigma, St. Louis, MO) was added to lower the pD of the solution to between 5 and 8. All 13C NMR spectra were recorded using a Bruker AM-500 spectrometer at 125.8 MHz using a 10-mm broadband probe. Composite pulse decoupling (CPD) in the inverse gated mode was used to decouple the 1H and 13C nuclei during data acquisition. A 10-s pulse delay was used between experiments to ensure complete relaxation of the 13C nuclei following the 90° excitation pulse. Because of the nature of the samples, 15,000, or in one case 7200 transients, were co-added to provide an adequate spectral signal-to-noise ratio. 4280
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Following data acquisition, the free induction decays (FIDs) were transferred to a Silicon Graphics workstation for data processing using Felix 97.0 (Biosym, San Diego, CA). The FIDs were zero-filled to 131,072 points, apodized using an exponential decay equivalent to 100-Hz line broadening, and Fourier transformed. The spectral baseline was corrected by fitting selected baseline points to a fifth-order polynomial. 13 C chemical shifts were measured relative to an external dioxane standard at 67.4 ppm. Functional group assignments of chemical shifts followed those of Thorn (22) and Leenheer (10): aliphatic carbons, 0-60 ppm; carbohydrate, ether, alcohol carbons, 60-90 ppm; aromatic plus olefinic carbons, 90-165 ppm; carboxyl, ester, amide carbons, 165-190 ppm; and ketone and quinone carbons, 190-220 ppm. Each spectrum was integrated from 0 to 220 ppm, and this area was normalized to 100%. Then integration was performed for the five spectral regions just defined to give the percentage of the total spectrum represented by each spectral region, and the carbon distributions were calculated for each of these regions for the Wakarusa River and Clinton Lake humic and fulvic acids. Additionally, aromaticity, fa, defined as the area under the spectral region from 110 to 165 ppm divided by the total area of the spectrum, was calculated for each sample (23). Aromaticity provides an indication of the aromatic character of the sample without taking into account the spectral region from 90 to 110 ppm where acetal, hemiacetal, and anomeric carbon resonances of the carbohydrates overlap with the aromatic carbon resonances (22, 23). Additionally, aggregate differences were used to compare the percentage distributions of integrated peak areas. Specifically, aggregate difference consists of the sum of the absolute values obtained from subtracting the respective percentage distributions from the two 13C NMR spectra being compared. Particular attention was given to those spectral regions that differed by (10% or more. Finally, integration of the 115-95 ppm region accounted for condensed tannins, which yield a characteristic peak at 105 ppm (24). Specifically, 20% of the integral between 90 and 60 ppm was subtracted from the 115-95 ppm integral to correct for anomeric carbon content (25), and then the resulting percentage was multiplied times %C to obtain the ratio of mg C/mg of sample. The δ13C determinations were made by reacting the lyophilized AHS with copper oxide in a sealed quartz tube (26) to produce carbon dioxide, which was subsequently analyzed for stable carbon isotopic composition using a continuous-flow isotope-ratio mass spectrometer. Isotopic results were reported in parts per thousand (‰) relative to the Vienna Pedee Belemnite (VPDB). Titrations (27) were performed to determine carboxylicacid and phenol contents of the Wakarusa River and Clinton Lake humic and fulvic acids. Determinations of LOP of the fulvic acids followed procedures adapted from Ertel et al. (28) and Orem et al. (29). Fulvic acids also were analyzed for C, H, O, N, and ash at Huffman Laboratories (Golden, CO) using methods detailed in Huffman and Stuber (30). Humic and fulvic acids were analyzed for C, H, N, and S using a LECO CHNS analyzer (Leco Corporation, St. Joseph, MI) and procedures described in Orem et al. (29). Results of elemental analyses were used to calculate atomic ratios: H:C, C:O, C:N, and C:S. FTIR spectra (ATI Mattston Genesis Series with Balston BFS-200 Air Purifier, Lexington, MA) were measured for KBr pellets prepared by incorporating 2 mg of AHS into 200 mg of optical grade KBr and compressing the pellet in a hydraulic press. Determinations of HAA5 and THM4 formation potentials were made on Wakarusa River and Clinton Lake humic and fulvic acids. Ten-mg (10-mg) masses of AHS samples were dissolved in organic free water, and the resulting solutions were dosed with a sufficient concentration of NaOCl to yield
FIGURE 3. 13C NMR spectra of Wakarusa River fulvic acid (top) and humic acid (bottom).
TABLE 2. Elemental Analyses (Ash Free), Atomic Ratios, Carboxylic Acid (RCOOH), and Phenol (PhOH) Contents, Lignin Oxidation Product Ratios, HAA5-THM4 Yields, and HAA5-THM4 Formation Potentials for Humic and Fulvic Acids Derived from Water Collected from the Wakarusa River and Clinton Lake in 12 Sample Collection Events between September 10 and October 10, 1996 FIGURE 2. Time series plots showing (A) precipitation, (B) elevation of Clinton Lake measured from the spillway house, and (C) concentrations of DOC in water collected from the Wakarusa River and Clinton Lake between September 10 and October 10, 1996.
TABLE 1. Concentrations of DOC and Organic Carbon Concentrations of AHS, Humic Acids, and Fulvic Acids Derived from Water Collected from the Wakarusa River and Clinton Lake in 12 Sample Collection Events between September 10 and October 10, 1996a parameter
Wakarusa River
Clinton Lake
mean DOC in mg/L coeff of variation median DOC in mg C/L composite DOC in mg/L % DOC as AHS AHS in mg/L humic acids in mg/L fulvic acids in mg/L humic/fulvic ratio
5.4 ( 0.7 13. 4.9 4.9 46 2.26 0.331 (0.665)b 1.93 (3.61) 0.172
4.1 ( 0.06 1.5 4.1 3.9 33 1.19 0.114 (0.282) 1.17 (2.18) 0.0974
a Organic matter concentrations are given in parentheses for humic and fulvic acids. b Organic matter concentrations obtained by dividing organic carbon concentrations by the percentage of elemental carbon.
a residual of 1.0 to 5.0 mg/L free chlorine at a pH of 7.0 maintained with phosphate buffer (31). After 7 days, the reactions between the free chlorine and the samples were quenched with sodium sulfite or ammonium chloride, and the samples were analyzed. Formation-potential experiments were performed each day because the NaOCl-dosing solution degraded over time. Assessing the quality of the NaOCl-dosing solution was accomplished by titrating a dilute solution of NaOCl with phenylarsine oxide using a Hach amperometric titrator (Hach Company, Loveland, CO). The endpoint was determined graphically and was used to calculate the free chlorine concentration in the NaOCl. The analysis of HAA5 and THM4 generated in the formation-potential experiments used ion-exchange liquidsolid extraction (32) and liquid-liquid extraction (33), respectively, and a Hewlett-Packard 5890 gas chromatograph with electron capture detection and a DB-5 column (J&W Scientific, Folsom, CA). Sample results were reported in micrograms per liter HAA5 and micrograms per liter THM4. HAA5 and THM4 formation potential values were calculated
Wakarusa River parameter elemental analyses %C %H %O %N %S atomic ratios H:C C:N C:S O:C titration data RCOOH in µeq/mg PhOH content in µeq/mg δ13C determinations (VPDB) lignin oxidation product ratios S/V C/V P/V Vac/al S ac/al P ac/al HAA5 yield in mg HAA5/mg sample HAA5 formation potential in mg/L THM4 yield in mg THM4/mg sample THM4 formation potential in mg/L a
Clinton Lake
humic acid
fulvic acid
humic acid
fulvic acid
49.70 4.44 nda 3.19 0.54
53.38 5.50 39.15 1.64 0.31
40.46 4.19 nda 4.35 0.75
53.71 5.85 38.36 1.69 0.38
1.06 18.2 246 nda
1.22 37.9 252 0.550
1.23 10.8 147 nda
1.29 37.0 375 0.536
3.7 2.1
6.1 1.7
nda nda
6.2 1.8
-26.0‰
-25.9‰
3.73
0.823 0.140 0.494 1.05 0.71 3.22 3.48
-26.8‰
-26.0‰
2.91
0.774 0.041 0.561 0.99 0.77 8.39 4.18
2.48
12.6
0.821
9.12
15.18
17.53
9.14
18.05
10.1
63.27
2.58
39.39
nd: not determined.
by dividing the HAA5 or THM4 bottle concentration by the 10-mg/L concentration of AHS, humic acid, or fulvic acid to obtain the yield (micrograms HAA5 or THM4/milligram of sample) and then multiplying the yield by the concentration of the humic component in the lake water (12). Additionally, multiplying the individual components of HAA5 and THM4 by their respective organic carbon percentages gave yields in mg C/mg of sample.
Results and Discussion DOC and AHS. Sporadic precipitation occurred between September 9 and October 10, 1996 (Figure 2). On September VOL. 34, NO. 20, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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TABLE 3. Integrated Peak Areas of 13C Nuclear Magnetic Resonance Spectra of Humic Acids and Fulvic Acids Derived from Water Collected from the Wakarusa River and Clinton Lake in 12 Sample Collection Events between September 10 and October 10, 1996 sample
component
Wakarusa River Clinton Lake Suwanee River Wakarusa River Clinton Lake Suwannee River Lake Fryxell
humic acid humic acid humic acid (22) fulvic acid fulvic acid fulvic acid (22) fulvic acid (35)
a
heteroaliphatic aliphatic total aromaticity ketone/quinone carboxylic acid aromatic 90-60 ppm 60-0 ppm 220-0 ppm (fa) 220-190 ppm 190-165 ppm 165-90 ppma 4 8 7 1 2 6 1
16 13 16 19 16 19 19
46 35 48 34 [3.0] 36 [4.4] 33 19
11 15 12 10 13 15 14
23 29 17 37 33 27 47
Percentages in braces: 20% of integrated area from 90 to 60 ppm subtracted from integrated area from 115 to 95 ppm.
25 and October 7, precipitation increased the elevation of Clinton Lake as well as the stage of the Wakarusa River. Prior to the precipitation events, DOC concentrations in the Wakarusa River were slightly greater than in Clinton Lake. Following the precipitation events, increased DOC concentrations in the Wakarusa River measured between September 29 and October 1 (6.5 to 7.0 mg C/L) elevated the mean DOC concentration (5.4 ( 0.7 mg C/L) higher than the median DOC concentration for the entire sample-collection period (4.9 mg C/L) (Table 1). However, median DOC concentrations closely approximated the DOC concentrations of the composite samples of water collected from the Wakarusa River and Clinton Lake. Therefore, the percentage of DOC as AHS was reported relative to the composite DOC concentrations rather than mean DOC concentrations. Overall, higher mean and median concentrations of DOC were determined in water collected from the Wakarusa River than in water from Clinton Lake between September 10 and October 10, 1996 (Figure 2). Accordingly, higher concentrations of humic and fulvic acids occurred in water collected from the Wakarusa River than in water from Clinton Lake and contributed to the recovery of nearly 1.9 g of fulvic acid from the Wakarusa River as opposed to 1.3 g of fulvic acid from Clinton Lake (Table 1). Thurman (2) noted that, for typical lake and river waters, fulvic acids occur in higher concentrations than humic acids, and the relative proportion of humic acids decreases from rivers to lakes. AHS constituted 46% of DOC in water collected from the Wakarusa River and approached the 50% value for typical nonblackwater rivers (2). AHS constituted 33% of the DOC in water collected from Clinton Lake and occurred within the 30 to 40% range as being typical for lakes (2). For lakes surrounded by forests, the proportion of DOC accountable as AHS can vary with the concentration of DOC. Mattson and Kortelainen (32) found that AHS constituted 40% of the DOC in lakes with concentrations of DOC less than 8 mg/L and as much as 57% in lakes with DOC concentrations greater than 8 mg/L. 13C NMR and Source Approximation. Distributions of integrated peak areas calculated from the 13C NMR spectra of AHS extracted from the Wakarusa River and Clinton Lake (Figure 3, Table 3) were compared to samples representing the Suwannee River in Georgia and Lake Fryxell in Antarctica (Table 4). The Suwannee River drains from the Okenfenokee Swamp and AHS dissolved in the Suwannee River originate from fresh terrestrial vegetation litter, leaf and root exuadates, and leaf leachates (35). AHS in the Antarctic Lake Fryxell originate from the degradation of particulate organic carbon derived from algae and bacteria in the sediments and lake water (36). The most notable contrast of the two environments is the lack of terrestrial vegetation in the Dry Valley region surrounding Lake Fryxell. AHS in lakes and reservoirs can originate from sources within (autochthonous bacteria and algae) and outside (allochthonous terrestrial vegetation). Thus, one can make comparisons of the Wakarusa River and 4282
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100 100 100 100 100 100 100 b
0.38 0.27 0.42 0.30 0.29 0.28 ndb
nd: not determined.
TABLE 4. Aggregate Differences Calculated from 13C NMR Spectral % Distributions and AHS Samples from Table 2a Wakarusa River Suwanne River HAb
Clinton Lake FAb Suwanne River FAb Lake Fryxell FAb
Humic Acid 12%
Fulvic Acid 13% 21% aliphatic +10% 29% Ar plus olefinic + 15% aliphatic -10%
Clinton Lake 32% Arc plus olefinic -13% aliphatic +12%
18% 36% Ar plus olefinic +17% aliphatic -14%
a Magnitudes and identifications of spectral regions given where differences are greater than or equal to 10%. b HA, humic acid; FA, fulvic acid. c Ar, aromatic.
Clinton Lake AHS samples to the Suwannee River and Lake Fryxell AHS samples to evaluate the contribution of allochthonous and allochthonous sources. Based on aggregate differences (Table 4), the Wakarusa River humic acid compared well with Suwannee River humic acid. However, Clinton Lake humic acid differed due to decreased aromatic plus olefinic and increased aliphatic contents. Overall, aggregate difference show that the Wakarusa River and Clinton Lake fulvic acids have the greatest similarity with a difference of 13%. The similarity between the Wakarusa River and Clinton Lake fulvic acids also is apparent with comparable aromaticities (fa (Table 3) and H:C ratios (Table 2)), O:C ratios, and carboxylic acid contents and phenol contents (Table 3). Clinton Lake fulvic acid also compared within 18% Suwannee River fulvic acid; whereas Wakarusa River fulvic acid differed due to increased aliphatic content. Both the Wakarusa River and Clinton Lake fulvic acids considerably differed from Lake Fryxell fulvic acid due to increased aromatic plus olefinic and decreased aliphatic contents. AHS derived from agal/bacterial sources high aliphatic and low aromatic contents (10, 36) where as the opposite holds true for AHS derived from terrestrial plants. Given this information, some preliminary conclusions can be made: (1) Wakarusa River humic acid and Clinton Lake fulvic acid probably originated from terrestrial vegetation; and (2) Wakarusa River fulvic acid may show the effect of bacterial/ algal inputs due to its increased aliphatic content. Finally, (3) Clinton Lake humic acid also may show the effects of bacterial/algal sources due to its decreased aromatic plus olefinic and increased aliphatic content relative to Suwannee River humic acid. Determination of Source. The effect of terrestrial sources on aquatic humic and fulvic acids from the Wakarusa River
FIGURE 4. Benzophenols and lignin phenols generated from CuO oxidation of aquatic fulvic acids from the Wakarusa River and Clinton Lake with example of tannin m-dihydroxybenzene chemical structure. and Clinton Lake became apparent after completion of δ13C determinations, titrations, and determination of LOP. Determination of δ13C ratios for both Wakarusa River and Clinton Lake aquatic humic and fulvic acids yielded values in the -26.8 to -25.9 ‰ VPDB range (Table 3). Such ratios occur within the range shown by terrestrial C3 vegetation to include most forest vegetation and cool-season grasses (37, 38). Phenol contents determined through titration of the fulvic acids were all > 0.5 µeq/mg (Table 3). Fulvic acids derived from terrestrial vegetation generally have phenol contents > 0.5 µeq/mg because of the contribution from lignin (36). Finally, fulvic acids from the Wakarusa River and Clinton Lake yielded sufficient quantities of LOP to be derived from terrestrial vegetation (Figure 4). Consideration of ratios of LOP allowed for more detail in the assignment of source vegetation (Table 3); S:V (syringyl:vanillyl) ratios in the 0.4to-2.8 range correspond to origination from nonwoody angiosperms (28, 39, 40), and P:V (parahydroxy:vanillyl) ratios in the 0.3-to-1.6 range denote derivation from deciduous leaves (40, 41). Thus, LOP results were consistent with the riparian gallery forest vegetation within the Wakarusa River Valley. However, elemental nitrogen contents and C:N ratios shown by the Wakarusa River and Clinton Lake humic and fulvic acids (Table 3) indicate the contribution of primary (algae) and secondary (bacterial) aquatic sources (10, 36). Additional evidence supporting aquatic sources includes the finding of H:C ratios > 1. Lacking aromatic components derived from tannins and lignins, fulvic acids derived from algal/bacterial sources in lakes are more aliphatic (10, 36). Fulvic acid in Clinton Lake yielded a carboxylic acid content in excess of 6.0 µg/eq. Such carboxylic acid contents are typical for lakes (2) and could indicate the effect of aquatic sources. The contribution of algal/bacterial sources is very apparent in the Clinton Lake humic acid because of its high
elemental nitrogen content (> 4%) and its lack of aromaticity. Humic acids normally are considered to be more aromatic than fulvic acids (2), but the H:C ratio for the Clinton Lake Humic acid closely corresponds-to-such ratios for the Wakarusa River and Clinton Lake fulvic acids (Table 3). Furthermore, Clinton Lake humic acid differs from Suwannee River humic acid with an aggregate difference of 32% due to decreased aromatic plus olefinic (-13%) and increased aromatic contents (+12%) (Table 4). The relatively high elemental N content of the Clinton Lake humic acid allowed the recognition of transmittance peaks associated with amides (42) (1662 cm-1 (amide I) and 1532 cm-1 (amide II)) in the FTIR spectra (Figure 5). The elemental nitrogen contents and C:N ratios of the Wakarusa River and Clinton Lake fulvic acids (Table 3) also fall within the range that indicate that AHS incorporate N from soil organic matter during the percolation of water through a soil profile. AHS obtained from leachates produced by soaking plant material in distilled water generally has elemental nitrogen contents less than 1% (13). Additionally, fulvic acids in water-supply reservoirs where vegetation is postulated to have washed in from the shores also displayed elemental nitrogen contents less than 1% (12). A similar origination from the direct leaching of water through vegetation without percolation through the soil profile can be postulated for the Suwannee River fulvic acid, which has an elemental nitrogen content of 0.56% and a C:N ratio of 107 (43). Nitrogen accumulates in the soil through numerous mechanisms such as adsorption onto clay and soil organic matter, precipitation of nitrogen-containing organic matter, and mineralization by heterotrophic bacteria (2). Classes of nitrogen-containing substances (proteinaceous materials, chlorophyll, nucleic acids, and alkaloids) enter the soil and become the source of N-heterocyclics that are identified in soil organic matter and AHS (44). AHS that incorporate soil VOL. 34, NO. 20, 2000 / ENVIRONMENTAL SCIENCE & TECHNOLOGY
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FIGURE 5. FTIR spectra of Wakarusa River humic acid, Wakarusa River fulvic acid, and Clinton Lake humic acid. organic matter as the result of the percolation of water through the soil profile generally yield C:N ratios less than 50 (13). The Wakarusa River and Clinton Lake AHS share similarities with soil interstitial water AHS on the basis of carboxylic acid contents, FTIR spectra, and H:C ratios. Carboxylic acid contents for the Wakarusa River and Clinton Lake fulvic acids are within respective ranges for fulvic acids in soil water beneath woody vegetation (5.7 to 6.3 µeq/mg) (13) and carboxylic acid contents in shallow soil solutions beneath forests (5.6 to 8.3 µeq/mg) (45). Overall, water collected from the Wakarusa River has a yellow coloration attributed to dissolved AHS. The FTIR spectrum for Wakarusa River humic acid showed a shelf at about 1653 cm-1 (Figure 5). Comparable peaks in the 1600- to 1650-cm-1 range were associated with humic acids extracted from yellow-colored soil water collected beneath woody vegetation (13). Finally, H:C ratios displayed by the Wakarusa River and Clinton Lake AHS compared favorably with the 1.1-to-1.4 range in H:C values for interstitial soil-water collected beneath grass and woody vegetation (13). However, H:C ratios could be a poor 4284
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indicator of the effects of interstitial soil-water because the typical range for H:C ratios in groundwater, rivers, and streams is from 1.1 to 1.2 (2). Increased H:C ratios are a better indicator of the effect of degradation as the loss of oxygen-containing functional groups during the degradation of AHS leads to decreased O:C ratios and increased aliphatic content (13). The O:C ratios of the Wakarusa River and Clinton Lake fulvic acids were less than 0.6 (Table 3) and indicate the loss of oxygen-containing functional groups, like alcohols, ethers, and carbohydrates. As a result, FTIR spectra for the fulvic acids (Figure 5) showed little detail in the 1030- to 1180-cm-1 range associated with alcohols, ethers, and carbohydrates (46, 47). Comparable lack of detail in this range also has been observed in the FTIR spectra of interstitial soil-water, groundwater, and streamwater fulvic acids (13), and water-supply reservoir fulvic acids (12). Additional evidence for degradation comes from the types and quantities of LOP derived from the Wakarusa River and Clinton Lake fulvic acids. Acid-to-aldehyde ratios of the LOP were greater than 0.7 (Table 3). Ertel et al. (28, 40) attributed such ratios to the oxidative degradation of lignin phenols in AHS. Overall, quantities of LOP, especially cinnamyl phenols, in fulvic acids decreased during transition from the Wakarusa River to Clinton Lake (Figure 4). Cinnamyl phenols are particularly susceptible to degradation (12). Several mechanisms can degrade AHS in a river-reservoir setting. Studies of AHS using proton NMR under WATR (water attenuation by transverse relaxation) conditions determined that the loss of oxygen-containing functional groups (carbohydrates, alcohols, ethers), deoxygenation of aromatic substituents, and increased alkyl content correlated with increasing soil depths and depths in the water column due to increased microbial diagenesis (48). Aquatic fungi produce enzymes that attack AHS yield products with reduced aromatic and increased aliphatic character (49). Compared to the 5.6 µeq/mg phenol content of fulvic acid leached from deciduous leaves (13), phenol contents of Wakarusa River and Clinton Lake fulvic acids are reduced (Table 3). The action of microbes and photodegradation has been shown to eliminate phenol chemical structures in AHS (50). Additionally, contribution of algally derived aliphatic organic matter to AHS in Clinton Lake also could reduce phenol contents. To summarize, AHS from the Wakarusa River and Clinton Lake show evidence of origination in riparian forest vegetation, reflect respective effects of soil organic matter and the contribution of organic matter from algal/bacterial sources, and bear evidence of biological degradation and photodegradation. AHS from the Wakarusa River and Clinton Lake fulvic acid bear more of the evidence of terrestrial vegetation. However, Clinton Lake humic acid with its high elemental N content displays more effects from algal and bacterial sources within the reservoir. HAA5 and THM4 Formation Potentials. Humic and fulvic acids from the Wakarusa River produced comparable yields of HAA5 and THM4 per unit mass, whereas fulvic acid from Clinton Lake yielded the most HAA5 and THM4 per unit mass (Table 3). Compared to the Wakarusa River, the higher yields for HAA5 and THM4 may imply that Clinton Lake contains greater numbers of precursor chemical structures capable of yielding DBP during chlorination. Wakarusa River fulvic acid also yielded larger amounts of 3,5-dihydroxybenzoic acid (3,5-DHBA) than Clinton Lake fulvic acid during lignin oxidation (Figure 4). 3,5-DHBA includes m-dihydroxybenzene (m-DHB) chemical structures capable of producing HAA5 and THM4 during chlorination (51-54). Although not specifically a product derived from lignin (55), the benzophenol 3,5-DHBA may also have a flavonoid origin (56). However on the basis of the quantities of carbon per sample mass, amounts of 3,5-DHBA generated by the lignin
FIGURE 6. Comparison of carbon yields for tannin m-dihydroxybenzenes, HAA5, THM4, and 3,5-dihydroxybenzoic acid for aquatic fulvic acids from the Wakarusa River and Clinton Lake.
combined THM4 formation potential for Clinton Lake slightly exceeded the proposed THM4 Stage II MCL. These results demonstrate that AHS containing tanninderived m-DHB chemical structures derived from sources outside and within the Wakarusa River and Clinton Lake can yield sufficient THM4 to approach regulatory levels. However, this finding comes with two qualifications. First, contributions of other nonhumic components of DOC containing precursor chemical structures are not addressed. From September 10 to October 10, 1996, AHS in the Wakarusa River and Clinton Lake accounted for only 46 and 33% of the DOC in these respective sources, even though the THM4 formation potential for Clinton Lake AHS could account for up to 70% of theTHM4 concentration in finished water at the Clinton Lake Water Treatment Plant. Second, the time of sample collection only represents part of one season: late summer before many leaves fell from deciduous trees. Over the course of a year, variable inputs such as leaf fall, runoff from rain, and snowmelt can affect DOC and AHS concentrations in the Wakarusa River and Clinton Lake. Seasonal variability and the variability of DOC (humic and nonhumic) ultimately affect the generation of disinfection byproducts during chlorination, and under less than ideal conditions, regulatory levels could be exceeded. Additional work should address the contribution of nonhumic precursors to disinfection byproduct formation potentials in the Wakarusa River and Clinton Lake, the identification what circumstances and what times of year and what conditions could result in exceeded regulatory levels. Armed with this information, as well as knowledge of potential chemical structures that yield DBPs, operators of water treatment plants can adjust treatment practices and protect public health by providing safer drinking water.
Acknowledgments oxidation of fulvic acids from both sources appear insufficient to produce the HAA5 and THM4 carbon yields found in this study (Figure 6). In contrast, amounts of carbon attributable to condensed tannin structures containing m-DHB chemical structures (105 ppm by 13C NMR (24)) occur in excess of these HAA5 and THM4 carbon yields. The relative abundance of tannin-derived m-DHB chemical structures relative to those derived from lignin also supports the contention that AHS are derived mainly from tannins rather than lignins (24, 25). Overall, the elevated HAA5 and THM4 yields for the Clinton Lake fulvic acid (Table 3) appear to correspond with the greater abundance of tannin-derived m-DHB chemical structures (Figure 6). The higher HAA5 and THM4 yields for the Clinton Lake fulvic acid did not carry through to greater formation potentials (Table 3). Humic and fulvic acids occurred in higher concentrations in the Wakarusa River than in Clinton Lake, so greater formation potentials are associated with the Wakarusa River. However, the combined (humic and fulvic) THM4 formation potential (Table 3: 42 µg/L) approach the 54.8 µg/L THM4 concentration for finished water from the Clinton Lake Water Treatment Plant determined on September 23, 1996 (55). Thus, Clinton Lake AHS, constituting 33% of the DOC, could account for as much as 70% of the THM4 concentrations determined after treating water from Clinton Lake. The U.S. EPA has proposed Stage I Maximum Contaminant Levels (MCL) of 60 µg/L for HAA5 and lowering the MCL for THM4 from 100 to 80 µg/L (14). Under Stage II MCLs, HAA5 would be reduced to 30 µg/L, and THM4 would be reduced to 40 µg/L. AHS from the Wakarusa River and Clinton Lake yielded HAA5 formation potentials that were less than the HAA5 Stage II MCL. However, the combined (humic and fulvic acids) THM4 formation potentials for the Wakarusa River approached the THM4 Stage I MCL, and the
This work was supported in part by National Science Foundation Grant CHE-9524514. The authors thank Drs. Andrew Borovik and Joseph Heppert (Department of Chemistry, University of Kansas) for use of their laboratory facilities during the FTIR work. The authors also thank Mr. L. Mike Pope, USGS, for asking crucial questions during his review, and Dr. Jerry A. Leenheer, USGS, for his review and directing our attention to the importance of the 95-115 ppm region of 13C NMR spectra. Finally, Dr. Shari Stamer, Water Quality Manager, City of Lawrence Department of Utilities, is thanked for her discussion of DBP concentrations at the Clinton Lake Water Treatment Plant. The use of brand, trade, or firm names in this article is for identification purposes only and does not constitute endorsement by the U.S. Geological Survey.
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Received for review December 14, 1999. Revised manuscript received July 11, 2000. Accepted July 17, 2000. ES991376J
