Sorption of Uranyl Cations onto Inactivated Cells of Alfalfa Biomass

W University Avenue, El Paso, Texas 79968-0513. Studies have shown that biomaterials have the capacity to adsorb heavy metals and metal oxo-cations fr...
0 downloads 0 Views 342KB Size
Download PDF
Environ. Sci. Technol. 2006, 40, 4181-4188

Sorption of Uranyl Cations onto Inactivated Cells of Alfalfa Biomass Investigated Using Chemical Modification, ICP-OES, and XAS J. G. PARSONS, K. J. TIEMANN, J. R. PERALTA-VIDEA, AND J. L. GARDEA-TORRESDEY* University of Texas at El Paso, Department of Chemistry, 500 W University Avenue, El Paso, Texas 79968-0513

Studies have shown that biomaterials have the capacity to adsorb heavy metals and metal oxo-cations from aqueous solution. In addition, previous studies have shown that biomaterials have the ability to bind uranyl cations from solution with capacities that are comparable to or greater than some commercially available synthetic ion-exchange resins. By using chemical modification, inductively coupled plasma optical emission spectroscopy (ICP-OES), and X-ray absorption spectroscopy (XAS), we have found that the primary functional group on alfalfa biomass responsible for the binding of uranyl cations from aqueous solution is the carboxyl functionality. Batch pH dependency experiments show a direct relationship between the increase in binding and the increase in pH (up to pH 4.5). XAS experiments showed that the major ligand involved in the binding of uranyl cations from aqueous solution was either a nitrogen or oxygen ligand with coordination numbers ranging from 6 to 10 ( 1.

Introduction Heavy-metal contamination of the aquatic environment has become an issue of importance with respect to environmental and human health. Special interests are raised when dealing with uranium contamination of the environment. Uranium is an element that is both highly toxic and radioactive. A major concern with uranium is its use in munitions and the possible subsequent contamination of the environment. Mobility and subsequent contamination of surface and groundwater are issues that are raised since uranium and the uranyl cations are highly soluble in the pH ranges of acid rain. While traditional methods used to remediate heavy-metal ions and uranium ions have been employed to clean up different contaminated sites, they have proven to be both costly and relatively ineffective. In addition, the use of traditional methods has side effects that include toxic byproducts from their production, usage and their inability to work in high salt concentrations (1). For example, ionexchange resins use toxic chemicals in their production that have to be treated before disposal, and some of these toxic chemicals are left in the polymeric matrix that may be mobilized during their use (1). Other methods such as coagulation or flocculation use toxic or hazardous chemicals directly in the remediation process to either complex or to * Corresponding author phone: (915) 747-5359; fax: (915) 7475748; e-mail: [email protected]. 10.1021/es060071j CCC: $33.50 Published on Web 05/26/2006

 2006 American Chemical Society

cause the metal ions to precipitate. These remediation techniques for heavy metals raise questions about worker health, safety, and environmental health. However, numerous research groups have investigated the use of biomaterials for the remediation of heavy metals. In addition, the use of live organisms, such as dandelions, lambs’ lettuce, and manganese-oxidizing bacteria, has also shown the ability of organisms to uptake and immobilize uranyl cations from solution (2, 3). It has been observed that, in plants, the uranyl cation exists in an immobilized phosphate from within/on the plant (2) However, in manganese-reducing bacteria the uranyl cation becomes incorporated into manganese oxides formed by the bacteria (3). The use of biomaterials has advantages over the aforementioned classical remediation techniques. It has been shown by different researchers that biomaterials are not fouled by high concentrations of hard cations, as are ion-exchange resins (4, 5). Furthermore, there are no toxic materials used in the production of biomaterials or released during their use. The binding of heavy metals to biomaterials has been well-documented in the literature (6-9). The biosorption of heavy metals such as copper(II), cadmium(II), chromium(III), and chromium(VI) has shown much promise with many different biosorbents such as alfalfa, sawdust, Datura innoxia, and different algae biomaterials (4, 8, 10, 11). Different direct or indirect techniques have been used to attempt to elucidate the binding sites of heavy metals to biomaterials (4, 8, 1012). For example Rayson et al. have used 113Cd NMR to elucidate the binding sites of cadmium in Datura innoxia with some success. The main functional group on the biomass responsible for heavy-metal binding has been shown to be the carboxyl groups (12). Kuyucak et al. have determined that carboxyl groups on inactivated cells of Ascophyllum nodosum are responsible for cobalt(II) binding (13). GardeaTorresdey et al., using different chemical modification techniques of biomaterials and analytical instrumental techniques, have shown that the functional groups responsible for heavy-metal binding are carboxyl groups and possibly amino functionalities on different biomaterials (1418). Biomaterials have also been investigated for their potential use in the remediation and/or recovery of uranium from aqueous solutions, high salt concentration solutions, acid mine drainage, and uranium process solutions (19-26). Yang and Volesky have investigated the interactions of uranyl cations with Sargassum seaweed, and they have modeled the interactions of proton exchange with uranyl cations (22). In addition they have investigated the effect of pH on biosorption, which showed a pH-dependent trend and an increase in binding with an increase in pH. Jansson-Charrier et al. investigated the sorption of uranium ions onto chitosan and found that it was effective in the treatment of leachates from mines (24). Psareva et al. have investigated the sorption of uranium onto cork biomass (25). Whereas, Liu et al. have studied the effects of cations and anions on the biosorption processes for uranium and found little or no effect from different cations and anions (26). Although studies on the biosorption of uranium have been performed, the investigation into the mechanism(s) of biosorption has not been studied to any great extent. This study on uranium biosorption was conducted using a variety of different chemical modification and analytical instrumental techniques. In addition, the focus of the source of uranium for the sorption studies was the uranyl cation [uranium(VI)], because of its solubility, stability, and mobility (27). Furthermore, the presence of uranium in environmental VOL. 40, NO. 13, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

4181

FIGURE 1. (A) pH profile of UO2(CH3COO)2 reacted with native and chemically modified alfalfa biomass between pH 2 and 4.5. (B) pH profile of UO2(NO3)2 reacted with native and chemically modified alfalfa biomass between pH 2.0 and 4.5. (C) pH profile of UO2(CH3COO)2 reacted with ion-exchange resins, and activated carbon between pH 2 and 4.5. (D) pH profile of UO2(NO3)2 reacted with resins, and activated carbon between pH 2 and 4.5. solutions and from mining waste is predominantly present as the uranyl cation (28-30). The alfalfa biomass was esterified to observe the effects of removing available carboxylic acid groups, which have been shown to be important in metal ion binding. The biomass was also amino modified in two different ways. The first amino modification was an acetylation reaction, to reduce the number of available amino ligands on the biomass for potential binding, and adding a carbonyl group to the biomass. The second amino modification to the alfalfa biomass was succination, which eliminates the amino groups from binding but at the same time adds additional carboxyl groups. The sulfhydryl groups on the biomass were eliminated to observe the effects of removing sulfhydryl groups from the biomass on uranium ion binding. Finally, the biomass was hydrolyzed, which was performed to observe the effect of increasing the number of carboxyl groups on the binding of the uranium ions to the alfalfa biomass. Additional studies were performed to further understand the binding process of uranium to alfalfa biomass using ion-exchange resins with well-defined functional groups. By using the known functional groups on ionexchange resins, an understanding of the functional groups involved in the binding process on the biomass can be achieved. In addition, the binding of uranium ions to activated carbon was investigated in order to observe the amount of uranium binding to carbon atoms on the biomass. Batch binding experiments were performed to investigate the effect of pH. Furthermore, all batch studies on the binding of the uranyl cations to the different sorbents were determined using ICP-OES (inductively coupled plasma optical emission spectroscopy). To identify the functional group(s) on the biomass responsible for metal ion binding, XAS (X-ray 4182

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 13, 2006

absorption spectroscopy) experiments were performed. XANES (X-ray absorption near edge structure) experiments were performed to investigate any changes in geometry and oxidation state of the uranium ions that could have occurred after binding. Finally, EXAFS (extended X-ray absorption fine structure) studies were performed to investigate the coordination numbers of the uranium bound to the alfalfa biomass and the interatomic bonding distances.

Methodology Alfalfa Biomass Collection. The Malone cultivar of alfalfa (Medicago sativa) was collected from controlled field studies at New Mexico State University, Las Cruces, NM. The alfalfa biomass was harvested and treated as previously reported (4). Chemical Modifications of Alfalfa Biomass: Esterification. The alfalfa biomass was esterified as per GardeaTorresdey et al., who have previously published an esterification method for alfalfa biomass (31, 32). This modification was performed by adding methyl ester functionalities to the alfalfa biomass to remove available carboxylic acid groups. After the reaction, the biomass was washed three times with deionized water and lyophilized. Hydrolysis. Hydrolysis of the alfalfa biomass was performed as previously published (32). This modification was performed to increase carboxylic acid functionalities to the biomass. This reaction was performed by reacting the biomass with concentrated sodium hydroxide for 1 h. After the reaction, the biomass was washed three times with deionized water and lyophilized. Sulfhydryl Modification. Modifications to the sulfhydryl and sulfur moieties on the alfalfa biomass were performed

FIGURE 2. (A) XANES of native and modified alfalfa biomass and activated carbon reacted at pH 2.0 with UO2(CH3COO)2. (B) XANES of native and modified alfalfa biomass at pH 2.0 with UO2(NO3)2. (C) XANES of native and modified alfalfa biomass and activated carbon reacted at pH 4.5 with UO2(CH3COO)2. (D) XANES of native and modified alfalfa biomass and activated carbon reacted at pH 4.5 with UO2(NO3)2. (E) XANES of model compounds UO2(NO3)2 and UO2(CH3COO)2 and the carboxyl resin reacted with the uranyl cations (with UO2(NO3)2)). as previously published (33). This modification was performed in order to remove available sulfur groups on the alfalfa biomass by changing sulfhydryl functionalities on the biomass to double bonded thiopyrdine functionalities. After the reaction, the biomass was washed three times with deionized water and lyophilized. Acetylation. The acetylation of the alfalfa biomass was performed as previously published (33). Carbonyl functionalities were added to the biomass to remove available amino groups on the alfalfa biomass. After the reaction was finished, the biomass was washed three times in deionized water and lyophilized. Succination. The biomass was succinated as previously published (33). To remove available amino groups

on the alfalfa biomass, carboxylic acid functionalities were added to the amino groups on the biomass. When the reaction had come to completion, the biomass was washed three times using deionized water and subsequently lyophilized. ICP-OES Sample Analysis. All ICP-OES analyses were performed using a Perkin-Elmer Optima 4300 DV spectrometer. The ICP-OES instrument was optimized before any studies were performed using a BEC (background equivalent concentrations) test. The recommended BEC value for uranium at 385.958 nm is 8.33 mg/L, which was obtained during this analysis. The parameters used for all uranium analyses were as follows: torch power of 1500 W; sample flow rate of 1.25 mL/min; nebulization rate of 0.65 L/min, VOL. 40, NO. 13, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

4183

FIGURE 3. Raw EXAFS of uranyl nitrate hexahydrate and uranyl acetate dihydrate reacted with native alfalfa biomass and chemically modified biomass at pH 2.5. with a 45 s read delay; and an integration time ranging between 10 and 20 s. pH Profiles. The pH profile studies were performed as previously published (31). A 400-mg sample of the native alfalfa biomass, chemically modified alfalfa biomass, carboxyl resin [Diaion WT01S (purchased from Supelco (Bellefonte, PA)], amino resin [Diaion CRB02 (purchased from Supelco (Bellefonte. PA)], and powdered activated carbon (purchased from Fisher Scientific) were weighed out. The weighed samples were washed three times, once with dilute nitric acid and twice with deionized water, to remove any soluble material that may interfere with the reactions. The biomass samples were centrifuged at 3000 rpm (Fisher Scientific 8K) for 5 min between each washing cycle and the supernatants were discarded. The washed samples were then diluted in 100 mL of deionized water and pH adjusted to the following pH values: 2.0, 3.0, 4.0, and 4.5. The pH was adjusted to pH 2.0 first using dilute nitric acid (0.01 M) and the pH was raised to the other pH values of interest using dilute sodium hydroxide (0.01 M). At each pH interval, a 4.0-mL aliquot of the sample solution was extracted in triplicate for statistical and quality control purposes. The extracted samples were then centrifuged again at 3000 rpm for 5 min and the supernatants were discarded. Solutions consisting of 1.0 mM of either UO2(NO3)2 or UO2(CH3COO)2 were pH adjusted and 4184

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 13, 2006

4.0-mL aliquots were added to the respective pH-adjusted (pH 2.0, 3.0, 4.0, and 4.5) adsorbent samples. In addition, at every pH value 4.0 mL aliquot control solution samples of the uranyl cations were extracted in triplicate and treated the same as the adsorbent samples for statistical and quality control reasons. The adsorbent samples and UO22+ cations were equilibrated for 1 h on a rocker. A 1.0 h reaction time was determined from previously determined studies on the time dependency of uranium binding to alfalfa biomass (34). After the 1-h equilibration period, the adsorbent samples and control solutions were centrifuged at 3000 rpm for 5 min and the supernatants were decanted and saved for ICP-OES analysis. All ICP-OES analyses were performed using a PerkinElmer Optima 4300 DV, and calibration coefficients of 0.98 or better were obtained for all calibrations. XAS Sample Preparation and Data Collection. Samples of 200 mg of ground 100 mesh (0.149 mm) alfalfa biomass (native or chemically modified), activated carbon, and the previously mentioned ion-exchange resins were washed three times once with dilute nitric acid and twice with deionized water. The washing was performed to remove any soluble material that may interact with the cation-biomass reactions. The samples were centrifuged at 3000 rpm for 5 min between each washing. The samples were then pH adjusted to their respective pH (either 2 or 4.5) and centrifuged at 3000 rpm. Subsequently to this, either 20 mL of pH-adjusted (2.0 or 4.5) 1000 ppm UO2(NO3)2 or UO2(CH3COO)2 was added to the samples. The sample and the UO22+ cations were allowed to react for 1 h and centrifuged again at 3000 rpm for 5 min. The supernatants were then decanted and the biomass, activated carbon, and ion-exchange resins were lyophilized. The lyophilization process entailed freezing the samples in liquid nitrogen for approximately 45 min until the samples were completely frozen. The frozen samples were then placed in a Labconco FreezeDry System (Freezone 4.5) until the water was removed from the samples. The uranium-laden samples were then packed into 1.0-mm sample plates, with Kapton tape windows for analysis at SSRL (Stanford Synchrotron Radiation Laboratory). The uranium LIII-edge XAS spectra were collected at SSRL on beam line 2-3 using a uranium oxide internal reference standard (LIII edge energy of 17.166 keV). The standard operating conditions of the beam line include energy of 3 GeV and current ranging from 60 to 100 mA. The samples were analyzed at room temperature using a Lytle detector to record the sample fluorescence. A Si (220 φ 90) double crystal monochromator was used for all measurements with a 1.0 mm slit. Model compounds UO2(NO3)2 and UO2(CH3COO)2 were diluted using boron nitride. The model compounds were diluted to give a one absorption unit change across the absorption edge. To reject higher order harmonics, the monochromator was detuned by 50% for all measurements. All spectra collected and analyzed were calibrated against the edge position of the uranium oxide foil. To improve the signal-to-noise ratios, several scans were averaged for each sample. XAS Data Analysis. The XAS data analysis of the samples and model compounds was performed using WinXAS and standard data reduction methods (35, 36). All the fluorescence spectra were calibrated according to the edge energy of the uranium oxide foil (17.166 keV). The background correction of the samples was performed using a one-degree polynomial fit to the pre-edge region. The XANES spectra were then extracted from 17.100 to 17.300 keV to investigate edge features. EXAFS spectra of the samples and the model compounds were extracted from the background-corrected XAS spectra, by converting the spectra into k-space (or wave vector space). The conversion into k-space was performed on the basis of the energy of the photoelectrons ejected from the sample,

FIGURE 4. (A) Raw EXAFS of uranyl nitrate hexahydrate model compound and uranyl nitrate reacted with the activated carbon, carboxylic acid resin, native alfalfa biomass, and chemically modified biomass. (B) Raw EXAFS of uranyl acetate dihydrate model compound and uranyl acetate reacted with the activated carbon, native alfalfa biomass, and chemically modified biomass. which was calculated from the first inflection point of the absorption edge. A spline of 7 knots of postedge region was then taken between 2.0 and 14.2 Å-1. By extracting the EXAFS to a k-space of 14.2 Å-1, the Debye-Waller factors and the noise in the spectrum were reduced. The EXAFS spectra were then k-weighted to 2 and Fourier transformed. The first- and second-shell EXAFS were extracted using WinXAS and backtransformed into k-space. The phases and amplitudes for the fittings were created using the ab initio code FEFF 8.00, and fittings were performed using the WinXAS software (36). The input files for the FEFF 8.00 software were created using ATOMS and crystallographic data of uranium compounds (38-40).

Results and Discussion The results of the effect of pH on the uranyl cation binding to the native and modified alfalfa biomass are shown in Figure 1 A and B. The general trend shown for both the UO2(NO3)2 and UO2(CH3COO)2 is an increase in binding with a corresponding increase in pH, which has been reported for other cations and biomaterials. The hydrolyzed and sulfhydrylmodified alfalfa biomass had the highest percentage of uranyl ion binding at 70 and 60%, respectively, whereas the acetylated and esterified alfalfa biomass had the lowest percentage of uranyl ion binding at approximately 30 to 40%. The data indicate that the carboxyl groups on the biomass are directly involved in the binding of the uranyl cation, due

to the large increase in percent metal bound after hydrolysis. The increase in binding observed after the sulfhydryl modification may be indicative of a surface charge interaction, indicating an indirect involvement of sulfur in the binding mechanism. The sulfur groups on the alfalfa biomass were neutral or positively charged in the pH ranges studied here. By blocking the sulfhydryl group, the charge is reduced or eliminated on the biomass, allowing the uranyl ions to get close to the biomass and bind. Similar binding trends to those obtained for the biomass were observed for the synthetic resins studied, as shown in Figure 1C,D. In Figure 1, parts C and D, the amino resin and the carboxyl resin have the highest percent uranyl cation binding at approximately 60 and 75%, respectively, and the activated carbon had the lowest percentage binding at approximately 20-30%. The data from the synthetic adsorbents indicates that both carboxyl and amino groups are possibly responsible for the binding of uranyl cations from aqueous solution. The data also indicates that carbon may also play a minor role in the binding of the uranyl cation from solution. The binding results of the amino resin in the pH profiles from the synthetic adsorbents indicates that amino groups are involved in the binding mechanism, as the pH binding trend closely resembles the alfalfa biomass binding trend. The carboxyl resin, however, has a large increase in binding at pH 3.0. The deviation from the trend by the carboxyl resin from the biomass may be explained by differences in the pKa VOL. 40, NO. 13, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

4185

TABLE 1. Edge Energies of the Chemically Modified, Uranium-Treated Biomass Samples at pH 2.0 and pH 4.5a uranyl acetate dihydrate

a

uranyl nitrate hexahydrate

sample

pH 2.0

pH 4.5

pH 2.0

pH 4.5

native alfalfa hydrolyzed alfalfa acetylated alfalfa sulfur-modified alfalfa activated carbon carboxyl resin

17.174 17.174 17.174 17.174 N/A N/A

17.173 17.174 17.173 17.174 17.173 N/A

17.174 17.173 17.173 17.174 NA 17.174

17.174 17.174 17.174 17.174 17.174 17.174

The edge energies of the uranyl nitrate hexahydrate and uranyl acetate hexahydrate were 17.176 and 17.175 keV, respectively.

of the carboxyl groups on the resin and the biomass. The increase in the binding after hydrolysis shows the importance of the carboxyl group in the binding mechanism. In addition, the acetylation modification showed that by removing amino groups there is not a large decrease in binding, approximately 10% at pH values of 4.0 and 4.5, compared to the native biomass. However, a decrease in the binding of the uranyl cation to the alfalfa biomass was observed at all pH values after acetylation. Similar decreases were observed with the esterified biomass in the pH profile studies. However, by replacing the amino group with a carboxyl group, the succination modification, the binding did not significantly change, indicating that there may not be many available amino groups on the biomass for binding. Results from the XAS studies are shown in Figures 2-4 and Tables 1-4 (Tables 2-4 are available as Supporting Information). The XANES studies shown in Figure 2 and Table 1, which show that the oxidation state of the uranyl cations does not change after reaction with the native or chemically modified alfalfa biomass or with the different synthetic resins studied. When studying LIII edges, the position of the white line indicates the oxidation state of the metal of interest. As shown in Figure 2 and Table 1, the position of the edge is not shifted between the model compounds and the samples, showing that the samples and the model compounds are at the same oxidation state. The shape of the XANES region, which includes the region from 20 eV before the inflection point of the edge to approximately 50 eV above the edge, provides information on the three-dimensional geometrical arrangements of the atoms around the central absorbing atoms (41). The XANES of the model compounds UO2(NO3)2 and UO2(CH3COO)2 have no observable differences, meaning that both compounds have the same geometrical arrangement. The XANES of the biomass samples and the adsorbents, however, do have one major observable difference from their parent model compounds. The white line feature is slightly less intense and broader, indicating that the bonding in the samples has changed somewhat in comparison to the parent model compounds. This broadening of the white line can be explained through steric hindrance and unequal bond lengths, since the other XANES features found in the model compound UO2(NO3)2 and UO2(CH3COO)2 around 17.176 and 17.175 keV are still present, but have less intensity in the sample spectra. A distortion in the geometry of a compound is shown in the XANES spectra through the dampening and broadening of spectral features of the XANES region (36). The EXAFS fittings are shown in Tables 2-4 (Supporting Information), and the raw EXAFS k-weighted to 2 are shown in Figures 2 and 3. All the fittings show the presence of the two axial oxygen ligands. The two axial oxygen ligands at approximately 1.80 Å have been reported by other authors for compounds with the UO22+ ion bound to different matrixes (2, 3, 42-44). The major difference in the EXAFS fittings of the samples is seen in the fittings of the equatorial oxygen atoms. Uranium(VI) cation, or the UO22+ ion, have coordination numbers of nearest neighboring atoms from 3 to 8 (45). The most stable coordination number is 8; however, a 4186

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 13, 2006

coordination number of 6 indicates an octahedral arrangement of atoms around the central uranium atom, which is common in ring-type structures (45). Ring type structures from the UO22+ cations have been observed with uranium nitrate in the presence of oxygenated organic ligands, where two nitrate ligands are still present (45). The EXAFS data obtained from this study show the presence of six to eight oxygen atoms in two different binding shells, as indicated by the difference in bond lengths (Tables 2-4, Supporting Information). Admittedly, there is noise in the EXAFS data, which could cause some error in the data interpretation at longer interatomic distances; thus, only the first two shells were fitted. The average coordination number in the equatorial binding shells is approximately six with the presence of two oxygen atoms in the axial shell, giving a total of eight coordinating oxygen atoms. This type of structure produces a system where there are two equally distant axial oxygen atoms and six equatorial oxygen atoms at mixed bond lengths. Two of the equatorial oxygen atoms are present from the adsorbent, with slightly varying bond lengths, and the other four equatorial oxygen atoms could be coordinated to water molecules, nitrate, or acetate ligands from the parent solution. This structure is suggested because of the XANES and EXAFS information. The loss of the intensity of some of the XANES features indicates that the uranium ions are still present in an 8-coordinate arrangement of atoms. Although somewhat distorted, the coordination numbers from the EXAFS are generally indicating a 6-8-coordination arrangement of the oxygen atoms from both the solution and the biomass. The data indicate that the biosorbents adsorb the UO22+ cations from solution through an ion-exchange process, as is observed with the ion-exchange carboxyl resin. This is supported by the change in percent adsorption with changing pH. The data also suggest that alfalfa biomass may be an inexpensive and viable means to remediate UO22+ from contaminated solutions.

Acknowledgments We acknowledge financial support from the National Institutes of Health (NIH) (Grant S06GM8012-33). We also acknowledge the financial support from the University of Texas at El Paso (UTEP), Center for Environmental Resource Management (CERM), through funding from the Office of Exploratory Research of the EPA (Cooperative Agreement CR-819849-01-4). The authors acknowledge the EPA Student Support Program through the Office of Air and Radiation. The authors also acknowledge the HCBU/OMI Environmental Technology Consortium, which is funded by the Department of Energy. Portions of this research were carried out at the Stanford Synchrotron Radiation Laboratory (SSRL), a national user facility operated by Stanford University on behalf of the US Department on Energy (DOE), Office of Basic Energy Sciences. The SSRL Structural Molecular Biology Program is supported by the DOE, Office of Biological and Environmental Research, and by the NIH, National Center for Research Resources, Biomedical Technology Program. Dr. Gardea-Torresdey acknowledges the Dudley family for

the Endowed Research Professorship in Chemistry. In addition, the authors would like to acknowledge the SSRL/ DOE funded Gateway Program and the Materials Corridor Initiative.

Supporting Information Available Results of FEFF 8.00 fittings of the EXAFS for the alfalfa biomass, activated carbon, and ion-exchange resins at pH 2.0 and 4.5 for reactions with uranyl nitrate and uranyl acetate, and results of FEFF 8.00 fittings of the EXAFS for the model compounds and the uranyl reacted with carboxyl resin at pH 4.5 and 2. This material is available free of charge via the Internet at http://pubs.acs.org.

Literature Cited (1) Gardea-Torresdey, J. L.; Tiemann, K. J.; Parsons, J. G.; Gamez, G.; Yacaman, M. J. Characterization of trace level Au(III) binding to alfalfa biomass (Medicago sativa) by GFAAS. Adv. Environ. Res. 2002, 6, 313-323. (2) Guenther, A.; Bernhard, G.; Geipel, G.; Reich, T.; Rossberg, A.; Nitsche, H. Uranium speciation in plants. Radiochim. Acta 2003, 91, 319-328. (3) Webb, S. M.; Fuller, C. C.; Tebo, B. M.; Bargar, J. R.. Determination of uranyl incorporation into biogenic manganese oxides using X-ray absorption spectroscopy and scattering. Environ. Sci. Technol. 2006, 40, 771-777. (4) Gardea-Torresdey, J. L.; Tiemann, K. J.; Parsons, J. G. Determination of trace level copper(II) binding to Medicago sativa (alfalfa) by graphite furnace atomic absorption spectroscopy with Zeeman background correction. Microchem. J. 2001, 69, 133-142. (5) Gardea-Torresdey, J. L.; Tiemann, K. J.; Gonzalez, J. H.; Rodriguez, O. Phytofiltration of hazardous metal ions by alfalfa: A study of calcium and magnesium. J. Hazard. Mater. 1997, 56, 169-179. (6) Lovley, D. R.; Coates, J. D. Bioremediation of metal contamination. Curr. Opin. Biotechnol. 1997, 8, 285-289. (7) Volesky, B.; Holan, Z. R. Biosorption of heavy metals. Biotechnol. Prog. 1995, 11, 235-50. (8) Martin-Dupont, F.; Gloaguen, V.; Granet, R.; Guilloton, M.; Morvan, H.; Krausz, P. Heavy metal adsorption by crude coniferous barks: A modelling study. J. Environ. Sci. Health, Pt. A: Toxic/Hazard. Subst. Environ. Eng. 2002, 37, 1063-73. (9) Churchill, S. A.; Walters, J. V.; Churchill, P. F.Sorption of heavy metals by prepared bacterial cell surfaces. J. Environ. Eng. 1995, 121, 706-11. (10) Srivastava, H. C. P.; Mathur, R. P.; Mehrotra, I. Removal of chromium from industrial effluents by adsorption on sawdust. Environ. Technol. Lett. 1986, 7, 55-63. (11) Holan, Z. R.; Volesky, B.; Prasetyo, I. Biosorption of cadmium by biomass of marine algae. Biotechnol. Bioeng. 1993, 41, 81925. (12) Ke, H. Y. D.; Rayson, G. D. Characterization of cadmium binding sites on Datura innoxia using cadmium-113 NMR spectrometry. Environ. Sci. Technol. 1992, 26, 1202-5. (13) Kuyucak, N.; Volesky, B. The mechanism of cobalt biosorption. Biotechnol. Bioeng. 1989, 33, 823-31. (14) Parsons, J. G.; Hejazi, M.; Tiemann, K. J.; Henning, J.; GardeaTorresdey, J. L. An XAS study of the binding of copper(II), zinc(II), chromium(III) and chromium(VI) to hops biomass. Microchem. J. 2002, 71, 211-219. (15) Tiemann, K. J.; Rascon, A. E.; Gamez, G.; Parsons, J. G.; Baig, T.; Cano-Aguilera, I.; Gardea-Torresdey, J. L.. Heavy metal binding by inactivated tissues of Solanum elaeagnifolium: Chemical and subsequent XAS studies. Microchem. J. 2002, 71, 133-141. (16) Gardea-Torresdey, J. L.; Arteaga, S.; Tiemann, K. J.; Chianelli, R.; Pingitore, N.; Mackay, W. Absorption of copper(II) by creosote bush (Larrea tridentata): Use of atomic and X-ray absorption spectroscopy. Environ. Toxicol. Chem. 2001, 20, 2572-9. (17) Gardea-Torresdey, J. L.; Becker-Hapak, M. K.; Hosea, J. M.; Darnall, D. W. Effect of chemical modification of algal carboxyl groups on metal ion binding. Environ. Sci. Technol. 1990, 24, 1372-8. (18) Tiemann, K. J.; Gardea-Torresdey, J. L.; Gamez, Gerardo; Dokken, K.; Sias, S.; Renner, M. W.; Furenlid, L. R. Use of X-ray Absorption

(19)

(20)

(21)

(22) (23) (24)

(25)

(26)

(27)

(28)

(29)

(30)

(31)

(32)

(33)

(34)

(35)

(36)

(37)

(38) (39) (40)

(41)

Spectroscopy and Esterification to Investigate Cr(III) and Ni(II) Ligands in Alfalfa Biomass. Environ. Sci. Technol. 1999, 33, 150154. Nakajima, A.; Tsuruta, T. Competitive biosorption of thorium and uranium by actinomycetes. J. Nucl. Sci. Technol. 2002, 528531. Bhainsa, K. C.; D’Souza, S. F. Uranium(VI) biosorption by dried roots of Eichhornia crassipes (water hyacinth). J. Environ. Sci. Health, Pt. A: Toxic/Hazard. Subst. Environ. Eng. 2001, A36, 1621-1631. Gonzalez-Munoz, M. T.; Merroun, M. L.; Omar, N. B.; Arias, J. M. Biosorption of uranium by Myxococcus xanthus. Int. Biodeterior. Biodegrad. 1997, 40, 107-114 Yang, J.; Volesky, B. Modeling uranium-proton ion exchange in biosorption. Environ. Sci. Technol. 1999, 33, 4079-4085. Popa, K.; Cecal, A. A review on biosorption of uranyl ions. Environ. Eng. Manage. J. 2003, 2, 69-75. Jansson-Charrier, M.; Guibal, E.; Roussy, J.; Surjous, R.; Le Cloirec, P. Dynamic removal of uranium by chitosan: Influence of operating parameters. Water Sci. Technol. 1996, 34, 169177. Psareva, T. S.; Zakutevskyy, O. I.; Chubar, N. I.; Strelko, V. V.; Shaposhnikova, T. O.; Carvalho, J. R.; Correia, M. J. N. Uranium sorption on cork biomass. Colloids Surf. A 2005, 252, 231-236. Liu, W.-J.; Xu, W.-C.; Wang, B.-E. Influence of cations and anion in solution on biosorption of uranium. Youkuangye 2004, 23, 143-146. Duff, M. C.; Coughlin, J. U.; Hunter, D. B. Uranium coprecipitation with iron oxide minerals. Geochim. Cosmochim. Acta 2002, 66, 3533-3547. Bernhard, G.; Geipel, G.; Brendler, V.; Nitsche, H. Uranium speciation in waters of different uranium mining areas. J. Alloys Compd. 1998, 271-273, 201-205. Mosselmans, J. F.; Bailey, E.; Schofield, P. A study of uranium speciation in acetate solutions at temperatures from 25 to 250 °C. J. Synchrotron Radiat. 2001, 8, 660-662. Abdelouas, A.; Lu, Y.; Lutze, W.; Nuttall, H. E.. Reduction of U(VI) to U(IV) by indigenous bacteria in contaminated ground water. J. Contam. Hydrol. 1998, 35, 217-233. Parsons, J. G.; Gardea-Torresdey, J. L.; Tiemann, K. J.; Gonzalez, J. H.; Peralta-Videa, J. R.; Gomez, E.; Herrera, I. Absorption and emission spectroscopic investigation of the phyto-extraction of europium(III) nitrate from aqueous solutions by alfalfa biomass. Microchem. J. 2002, 71, 175-183. Tiemann, K. J.; Gamez, G.; Dokken, K.; Parsons, J. G.; GardeaTorresdey, J. L. Chemical modification and X-ray absorption studies for lead(II) binding by Medicago sativa (alfalfa) biomass. Microchem. J. 2002, 71, 287-293. Gardea-Torresdey, J. L.; Tiemann, K. J.; Dokken, K.; Gamez, G. Investigation of metal binding in alfalfa biomass through chemical modification of amino and sulfhydryl ligands. Proc. Conf. Hazard. Waste Res. 1998, 111-121. Parsons, J. G.; Tiemann, K. J.; Peralta, V.; LeMone, D. V.; GardeaTorresdey, J. L. Phytroremediation generalized overview with specific focus on the remediation of uranium from water. Proceedings of the Waste Management Conference; February 2004, Tucson, AZ. Ressler, T.. WinXAS: A program for X-ray absorption spectroscopy data analysis under MS-Windows. J. Synchrotron Radiat. 1998, 5, 118-122. Parsons, J. G.; Aldrich, M. V.; Gardea-Torresdey, J. L. Environmental and biological applications of extended X-ray absorption fine structure (EXAFS) and X-ray absorption near edge structure (XANES) spectroscopies. Appl. Spectrosc. Rev. 2002, 37, 187222. Ankudinov, A. L.; Ravel, B.; Rehr, J. J.; Conradson, S. D. Realspace multiple-scattering calculation and interpretation of X-ray-absorption near-edge structure. Phys. Rev. B 1998, 58, 7565-7566. Ravel, B. ATOMS: Crystallography for the X-ray absorption spectroscopist. J. Synchrotron Radiat. 2001, 8, 314-316. Fleming, J. E.; Lynton, H. Crystal structure of uranyl nitrate hexahydrate. Chem. Ind. 1960, 1416-17. Howatson, J.; Grev, D. M.; Morosin, B. Crystal and molecular structure of uranyl acetate dihydrate. J. Inorg. Nucl. Chem. 1975, 37 (9), 1933-5. Koningsberger, D. C.; Prins, R.; Editors. Chemical Analysis, Vol. 91: X-ray Absorption: Principles, Applications, Techniques of EXAFS SEXAFS, and XANES; John Wiley & Sons: New York, 1988; p 673.

VOL. 40, NO. 13, 2006 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

9

4187

(42) Pompe, S.; Schmeide, K.; Bubner, M.; Geipel, G.; Heise, K. H.; Bernhard, G.; Nitsche, H. Investigation of humic acid complexation behavior with uranyl ions using modified synthetic and natural humic acids. Radiochim. Acta 2000, 88, 553-558. (43) Allen, P. G.; Shuh, D. K.; Bucher, J. J.; Edelstein, N. M.; Reich, T.; Denecke, M. A.; Nitsche, H. EXAFS determinations of uranium structures: The uranyl ion complexed with tartaric, citric, and malic acids. Inorg. Chem. 1996, 35, 784-7. (44) Allen, P. G.; Shuh, D. K.; Bucher, J. J.; Edelstein, N. M.; Palmer, C. E. A.; Silva, R. J.; Nguyen, S. N.; Marquez, L. N.; Hudson, E. A. Determinations of uranium structures of EXAFS: Schoepite

4188

9

ENVIRONMENTAL SCIENCE & TECHNOLOGY / VOL. 40, NO. 13, 2006

and other U(VI) oxide precipitates. Radiochim. Acta 1996, 75, 47-53. (45) Cotton, F. A.; Wilkinson, G.; Bochmann, M.; Murillo, C. Advanced Inorganic Chemistry, 6th ed.; Wiley: New York, 1998; 1248 pp.

Received for review January 11, 2006. Revised manuscript received April 14, 2006. Accepted April 24, 2006. ES060071J