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a Stratum Engineering, Inc., 3751 Pennridge Dr., Suite 119, Bridgeton, MO 63044
b Dep. of Plant and Soil Sciences, 052 Agricultural Hall, Oklahoma State Univ., Stillwater, OK 74078
c Veterinary Medicine Diagnostic Laboratory, 1600 E. Rollins St., Univ. of Missouri-Columbia, Columbia, MO 65205
* Corresponding author (bastan{at}okstate.edu)
Received for publication February 17, 2002.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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hydroxylamine hydrochloride (51.7%) > phosphate (10.5%), acetate (7.16%) > water (0.15%). The strongest relationship between arsenic determined by soil chemical extraction and in vivo bioavailable arsenic was found for hydroxylamine hydrochloride extractant (r = 0.88, significant at the 0.01 probability level). Comparison of the amount of arsenic extracted by soil methods with bioavailable arsenic showed the following trend: ammonium oxalate, hydroxylamine hydrochloride > in vivo > phosphate, acetate > water. The amount of arsenic dissolved in the stomach (potentially bioavailable) is between surficially adsorbed (extracted by phosphate or acetate) and surficially adsorbed + nonsurficial forms in Fe and Mn oxides (extracted by hydroxylamine hydrochloride or ammonium oxalate). Soil extraction methods that dissolve some of the amorphous Fe, such as hydroxylamine hydrochloride, can be designed to provide closer estimates of bioavailable arsenic.
Abbreviations: ABA, absolute bioavailability • HG, hydride generation • ICP, inductively coupled plasma • RBA, relative bioavailability • UEF, urinary excretion factor
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Incidental soil ingestion by children is an important exposure pathway in assessing public health risks associated with exposure to arsenic-contaminated soils (Dudka and Miller, 1999). The importance of soil ingestion by children as a health issue has been reported by numerous researchers and fully illustrates the importance of this pathway in terms of subsequent chemical exposure (Calabrese et al., 1989; Davis et al., 1990).
Most risk from arsenic is associated with the forms of arsenic that are biologically available for absorption, or "bioavailable" to humans. A bioavailable chemical is the portion of a chemical dose that enters the systemic circulation from an administered dose. Presently, methods are not available to quantify the amount of bioavailable arsenic in soils to more accurately assess risk from incidental ingestion of arsenic-contaminated media. The method routinely used to characterize arsenic in soils and solid wastes at hazardous waste sites for remedial investigation and risk assessment purposes is by hot acid extraction using USEPA SW-846, Method 3050 (USEPA, 1986). However, total content may not be related to solubility or bioavailability. Therefore, the use of total arsenic content to quantify daily chemical intake during exposure assessment is unlikely to provide an accurate assessment of risk from contaminated media.
The bioavailability of metals, especially lead and arsenic, in some mining wastes has been assessed by conducting expensive and lengthy dosing trials using animal models. Immature pigs have therefore been used successfully as a model for gastrointestinal function of children (Miller and Ullrey, 1987; Weis and LaVelle, 1991). To overcome the difficulties and expenses associated with the conduct of animal dosing trials to assess bioavailability of metals in soils, research efforts have been directed toward the development of chemical methods. Rodriguez et al. (1999) and Ruby et al. (1996) evaluated the effectiveness of using chemical extracts that simulate the gastrointestinal environment as a means of quantifying bioavailable metals with respect to the soil ingestion pathway. Methods to simulate the gastrointestinal environment, in vitro methods, have been recently reviewed (Basta et al., 2001a; Ruby et al., 1999). Another approach to evaluate solubility and availability of chemical elements in soils involves the use of selective chemical extractants. Selective extractants have been historically used to evaluate plant nutrients in soil. Work performed as early as the 1930s by soil scientists (e.g., Bray, Hester, Morgan, Spurway, Truog) has demonstrated the need to measure "labile" pools rather than total content of nutrients to evaluate conditions for optimum plant growth (Peck and Soltanpour, 1990). Chemical extraction procedures for a limited number of heavy metal contaminants have been correlated with plant uptake (Adriano, 2001; Basta and Gradwohl, 2000; Soon and Bates, 1982; Xian, 1989). To our knowledge, the ability of soil chemical extraction methods to measure arsenic bioavailability to humans via the soil ingestion pathway has not been reported.
Chemical fractionation methods, based on sequential extraction procedures, have been used to determine the amount of contaminant in specific chemical pools (Pierzynski, 1998). Most chemical fractionation methods have been used to measure heavy metals in ion-exchangeable, surficially adsorbed, precipitated, organic chelated, and occluded chemical pools in baseline soils (Tessier et al., 1979); in sewage sludge–amended soils (Sposito et al., 1982); and in contaminated soils (Gibson and Farmer, 1986; Hickey and Kittrick, 1984; Ma and Rao, 1997). Both heavy metal solubility and bioavailability decrease with each successive step of the sequential extraction.
Because arsenic is chemically similar to phosphorus, it has been evaluated by using chemical extractants developed to measure the various pools of phosphate. Researchers have used this approach to specifically look at various pools of arsenic in soils (Gruebel et al., 1988; Johnson and Barnard, 1979; McLaren et al., 1998). A two-step sequential extraction method has been used (Chao and Zhou, 1983; Shuman, 1982) to determine arsenic associated with amorphous iron oxides, manganese oxides, and/or organic matter in wastes, but an inherent problem associated with this method is the readsorption of arsenic to soil residue after dissolution of iron oxides (Gruebel et al., 1988). Amacher and Kotuby-Amacher (1994) incorporated phosphoric acid into acid hydroxylamine hydrochloride extraction of soils and mine spoils to prevent readsorption of arsenic onto ferrihydrite minerals. Incorporating 0.1 M phosphate into acid hydroxylamine hydrochloride extraction of soils reduced arsenic readsorption by goethite from 100 to 0% (Jackson and Miller, 2000). The ability of soil chemical extraction methods to measure arsenic phytoavailability has been reviewed recently by Adriano (2001). However, the ability of soil extraction methods to assess arsenic bioavailability to humans via the soil ingestion pathway has not been reported.
A chemical extractant procedure will provide a rapid, inexpensive testing method to obtain scientifically derived data to select appropriate remedies at contaminated sites that are cost effective and protective of human health and the environment. A measure of bioavailable arsenic will also serve to lower the uncertainty surrounding the quantification of potential risks arising from exposure to arsenic-contaminated media. The objective of our study is to evaluate the ability of previously reported chemical extractants to measure the bioavailable fraction of arsenic in contaminated soil and solid wastes as measured in vivo using immature swine dosing trials.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Immature Swine Dosing Trial
Standard operating procedures developed by Dr. Stan Casteel at the University of Missouri-Columbia Veterinary Medical Diagnostic Laboratory, approved by the USEPA Region 8 (Casteel, 1995), were used in the immature swine trials. Intact male swine weighing 10 to 12 kg were randomly assigned to treatment groups consisting of a calcine dosing group, slag dosing group, negative control group (no soil), and positive control group (oral Na2AsO4). Five swine were used per treatment group, with the exception of three swine per negative control group. All swine were individually housed in arsenic-free cages and fed water and a grower ration formulated by the University of Missouri feed mill. After a 3-d acclimation period, the swine were exposed to soil and treatment doses. Swine were dosed with 6.25 mg of soil per kg body weight per day with one-half of the dose administered 2 h before feeding in the morning and the remaining half given 2 h before the afternoon feeding. Soil doses were placed in the center of a 5-g portion of moistened low-arsenic and low-lead diet material (Ziegler Brothers, Gardners, PA) and hand-administered to each animal. The soil mass per dose varied from 23 to 30 mg initially and was adjusted every 3 d to account for growth of the swine. Soil masses delivered during the final 3 d of the dosing trial ranged from 41 to 45 mg. Every 3 d thereafter, for five collection periods, 24-h excretions of urine were collected from each swine. The urine samples were collected in a stainless steel pan fitted with a nylon screen to minimize contamination with feces, spilled food, or other debris. Plastic diverters were used to minimize urine dilution with drinking water spilled by the animals from the watering nozzle into the collection pan. Collection containers were emptied twice daily into a separate holding container to ensure no loss because of overflow. The urine samples were filtered (Whatman [Maidstone, UK] #2), placed into plastic bottles, and preserved to pH 2 with concentrated HCl. Urinary samples were packed securely in coolers on ice and shipped by overnight carrier under chain-of-custody procedures to Oklahoma State University for arsenic analysis. Following an additional filtering through 0.45-µm filters, arsenic analysis was performed by a Thermo Jarrell Ash IRIS high resolution inductively coupled plasma (ICP) with hydride generation (HG) (Thermo Elemental, Franklin, MA). Arsenic was determined at an analytical wavelength of 189.042 nm. Spectral scans and other quality control procedures showed this wavelength was free of spectral interferences commonly observed on ICP instruments with poorer resolution (i.e., Fe, Al, and Ca interferences).
Urine samples were analyzed consistent with standard operating procedures developed by Dr. Stan Casteel at the University of Missouri-Columbia Veterinary Medical Diagnostic Laboratory, approved by USEPA Region 8, for analysis of arsenic in urine collected from immature swine dosing trials. Urine samples were prepared for analysis as follows. Urine (25 mL) was transferred to an acid-washed, 100-mL beaker. Methanol (3.0 mL), five drops of antifoam agent, 10.0 mL of 40% (w/v) magnesium nitrate hexahydrate, and 10.0 mL of concentrated trace metal–grade nitric acid were added to the urine sample. The beaker was covered with a watch glass and placed on a hot plate to reflux for 8 to 12 h at 70 to 80°C. Subsequently, the temperature was increased to 200°C after removal of the watch glass to allow faster evaporation and heated to complete dryness (8–12 h). After cooling, dried samples in beakers were transferred to a cool muffle furnace and ignited according to the following temperature program: ramp to 500°C at 1°C/min, then held at 500°C for 3 h, turned off, and allowed to cool. Deionized water (5 mL) and concentrated trace metal–grade hydrochloric acid (5 mL) was added to the ignited residue and heated until the residue dissolved. The sample digest was diluted with deionized water to 50 mL.
Samples were mixed with a solution of 10% HCl, 10% KI, and 5% ascorbic acid for at least 16 h before analysis of arsenic by ICP–HG. Hydride generation was achieved by automated mixing of 2.2 mL min-1 1% NaBH4 in 0.1 M NaOH solution with treated sample solutions delivered at a flow rate of 2.2 mL min-1.
Quality control of urine analyses included the use of Na9tional Institute of Science and Technology (NIST) Standard Reference Material (SRM) 2670, Toxic Metals in Freeze-Dried Urine. This standard was included in each group of urine samples during urine dry ashing and wet digestion before ICP analysis. Ten replicate analyses of SRM 2670 showed our procedures measured 102.9 to 107.5% with a mean of 103.8% of the certified arsenic concentration. Analytical procedures were very precise with a coefficient of variation of 1.23% for the above analyses.
In Vivo Arsenic Bioavailability Calculations
The amount of arsenic absorbed through the gastrointestinal tract (bioavailable arsenic) may be described in absolute or relative terms. Absolute bioavailability (ABA) is the ratio of the amount of arsenic absorbed compared with the amount ingested, as in the following equation:
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Relative bioavailability (RBA) is the ratio of the absolute bioavailability of arsenic present in test material (study soil) compared with the absolute bioavailability of arsenic in an appropriate (e.g., high solubility in water) reference material, as in the following equation:
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In our study, we selected the Na2AsO4·7H2O reference material as the control because it is a readily soluble form of arsenic that is easily absorbed and because the majority of arsenic in our contaminated soils and solid media occurred as arsenate. Arsenic excretion in urine was found to be a linear function of the administered dose and was approximately independent of time after 5 d of exposure during dosing trials. In most animals, including pigs, absorbed arsenic is excreted primarily in urine. Thus, the urinary excretion fraction (UEF), defined as the amount excreted in the urine divided by the amount dosed, is a reasonable approximation of the oral absorption fraction, or ABA. However, this ratio will underestimate total absorption because some absorbed arsenic is excreted in the feces via the bile and some enters tissue compartments (e.g., liver, kidney, skin, hair, etc.) from which it is cleared very slowly or not at all. Thus, the urinary excretion fraction is not equated with the ABA. The UEF can be used, however, to compute the RBA as follows:
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All in vivo bioavailabilities of arsenic in this study are reported as arsenic relative bioavailability.
Chemical Fractionation Procedures
Soils were extracted by five soil chemical extraction methods. The various extracting solutions and conditions for each method are described below. All samples were extracted in triplicate and included appropriate reagent blanks and spikes.
Deionized Water Extraction (Huang and Fujii, 1996)
Soil (1 g of <250 µm) was placed into a 50-mL polycarbonate centrifuge tube and mixed with 20 mL of deionized water. The tube was sealed using a neoprene stopper and placed in a horizontal position on a reciprocal laboratory shaker. The tubes were shaken for 1 h and then centrifuged at 8000 rpm for 5 min. The supernatants were filtered through a 0.45-µm membrane filter, and the filtrate was acidified to pH < 2 with concentrated HCl for preservation before arsenic analysis. Arsenic analysis was performed by ICP–HG. To prepare the filtered soil extracts for ICP–HG analysis, a 10.0-mL aliquot of soil extract was placed into a test tube and mixed with 3.3 mL concentrated HCl and 4.0 mL of a solution containing 10% potassium iodide and 1% ascorbic acid. After a reaction period of at least 1 h, arsenic was determined by ICP–HG as described previously under urine analysis used for in vivo determination of bioavailable arsenic.
Sodium Acetate Extraction (Tessier et al., 1979)
Soil (1 g of <250 µm) was placed into a 50-mL polycarbonate centrifuge tube and mixed with 20 mL of 1.0 M sodium acetate solution adjusted to pH 5.0. The tube was sealed using a neoprene stopper and placed in a horizontal position on a reciprocal laboratory shaker. The tubes were shaken for 1 h and then centrifuged at 8000 rpm for 5 min. The supernatants were filtered through a 0.45-µm membrane filter and acidified to pH < 2 with concentrated HCl. Arsenic analysis was performed by ICP–HG as previously described.
Phosphate Extraction (Yamamoto, 1975)
Soil (1 g of <250 µm) was placed into a 50-mL polycarbonate centrifuge tube and mixed with 20 mL of a solution consisting of 3:2 (v/v) 0.1 M Na2HPO4 and 0.1 M NaH2PO4 (Yamamoto, 1975). The tube was sealed using a neoprene stopper and placed in a horizontal position on a reciprocal laboratory shaker. The tubes were shaken for 8 h and then centrifuged at 8000 rpm for 5 min. The supernatants were filtered through a 0.45-µm membrane filter and acidified to pH < 2 with concentrated HCl. Arsenic analysis was performed by ICP–HG as previously described.
Hydroxylamine Hydrochloride Extraction (Chao and Zhou, 1983 modified by Amacher and Kotuby-Amacher, 1994)
Soil (1 g of <250 µm) was placed into a 250-mL polystyrene centrifuge bottle and mixed with 250 mL of a solution containing 0.25 M NH2OH·HCl, 0.25 M HCl, and 0.025 M H3PO4 (Amacher and Kotuby-Amacher, 1994). The centrifuge bottles were placed into a 70°C water bath and shaken for 2 h. The bottles were then centrifuged for 10 min at 6000 rpm, and the supernatants were filtered through a 0.45-µm membrane filter. Arsenic analysis was performed by ICP.
Ammonium Oxalate Extraction (Shuman, 1982)
Soil (1 g of <250 µm) was placed into a 250-mL polystyrene centrifuge bottle and mixed with 50 mL of a solution containing 0.2 M (NH4)2C2O4, 0.2 M C2H2O4, and 0.1 M ascorbic acid. The oxalate-extracting solution pH was adjusted to 3.0 (Shuman, 1982). The centrifuge bottles were placed into a 100°C water bath and shaken for 15 min. The bottles were then removed and filtered through a 0.45-µm membrane filter. The solid material on the filter was then washed with an additional 50 mL of the fresh extracting solution to yield a final volume of 100 mL. Arsenic analysis was performed by ICP.
Determination of Total Arsenic Content
Total arsenic was determined by hot acid digestion with HNO3 and H2O2 according to USEPA SW-846, Method 3050 (USEPA, 1986). Arsenic analysis was performed by ICP.
Chemically Extracted Arsenic Bioavailability Calculations
The standard analysis for soil metal content, including arsenic, during the investigation of the nature and extent of contamination of hazardous waste sites is by hot digestion with HNO3 and H2O2 according to USEPA SW-846, Method 3050 (USEPA, 1986). The resulting total metal concentration is then used for estimating risks to human health. The realization that probably not all (<100%) of the total metal measured by complete digestion is bioavailable has led risk assessors to use a fraction (percentage) of total metal that better represents the fraction that is bioavailable in the risk calculation. For our chemically extracted results, bioavailable arsenic is calculated by dividing the arsenic concentration measured by the various chemical speciation extractions by that measured as total arsenic (all on a concentration in soil basis), as described by the following equation:
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Statistical Methods
Analysis of variance using a randomized complete block design and subsequent separation of means by Duncan's multiple range test (SAS Institute, 1988) was used to compare results between soil chemical extraction and in vivo methods. Linear regression was used to determine agreement between soil chemical extraction and in vivo results. Linear regression parameters (slope = 1 and intercept = 0) were evaluated using t tests to determine agreement between methods.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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Chemically Extracted Relative Bioavailability of Arsenic
Results of the various arsenic soil chemical extractants were compared with bioavailable arsenic values obtained from immature swine dosing trials. Because these extractions were conducted separately rather than performed sequentially, more than one chemical fraction of arsenic pool was extracted. Very little arsenic was extracted by the deionized water, generally <5 mg kg-1 of water-soluble arsenic (Table 3). The sodium acetate extractant, which measures the weakly adsorbed pool of arsenic as well as the water-soluble pool, extracted on average 50 times more arsenic than water (Table 3). The phosphate solution extracted a greater portion of arsenic than the sodium arsenate solution; results ranged between 70 and 381 mg kg-1 of arsenic. The phosphate-extracted fraction represents some of the strongly (specifically) adsorbed fraction of arsenic as well as the water-soluble and weakly adsorbed fractions. All of three solutions (water, sodium acetate, and phosphate) extracted adsorbed arsenic to the surface of soil particles.
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hydroxylamine hydrochloride (41.8%) > phosphate (5.45%), acetate (2.98%) > water (0.041%). This trend suggests that most of the arsenic in these soils is inside the soil mineral matrix and can only be released when iron oxides and other minerals that occlude the arsenic are dissolved by the chemical extractant (i.e., ammonium oxalate and hydroxylamine hydrochloride). Some extractants approach 100% of total arsenic, such as the hydroxylamine hydrochloride or ammonium oxalate extractant for some of the study samples. Much lesser amounts of surficially adsorbed arsenic were extracted by phosphate and acetate than reagents that dissolved mineral forms occluding arsenic (ammonium oxalate, hydroxylamine hydrochloride). Very little of the arsenic was extracted with water (Table 3). Chemical extraction results presented here demonstrate that the amount of arsenic in chemical pools was: inside mineral matrix > surficially adsorbed > water soluble. The relationship between arsenic extracted by soil chemical methods and in vivo bioavailable arsenic was determined by linear regression (Fig. 2) . Three chemical extractant methods that showed significant linear regression coefficients, r values, were water (Fig. 2A, r = 0.68, significant at the 0.01 probability level), sodium acetate (Fig. 2B, r = 0.57, significant at the 0.05 probability level), and hydroxylamine hydrochloride (Fig. 2D, r = 0.88, significant at the 0.01 probability level). The strongest relationship between arsenic measured by a chemical extraction method and in vivo bioavailable arsenic was found for hydroxylamine hydrochloride.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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ACKNOWLEDGMENTS |
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REFERENCES |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONCONCLUSIONSREFERENCES |
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