STUDY OF THE LIGANDS INVOLVED IN METAL BINDING TO ALFALFA BIOMASS
K.J. Tiemann1, J.L. Gardea-Torresdey1, S. Sias1, G. Gamez1, O. Rodriguez1, M.W. Renner2, and L.R. Furenlid2
1Department of Chemistry, The University of Texas at El Paso, El Paso, Texas 79968, (915) 747-5359; 2Department of Applied Science, National Synchrotron Light Source, Brookhaven National Laboratory, Upton, NY 11973-5000
Previously performed studies have shown that the alfalfa shoot biomass can bind an appreciable amount of copper(II), nickel(II), cadmium(II), chromium(III), lead(II), and zinc(II) ions from aqueous solution. Of the seven different alfalfa populations studied, Malone and African demonstrated the highest capacity for metal binding. Laboratory experiments were performed to determine the pH profiles, time dependency, capacity for metal binding, as well as the recovery of the metals bound. For most of the metal ions studied, the biomass showed a high affinity for metal binding around pH 5.0 within a short time period. Binding capacity experiments revealed the following amounts of metal ions bound per gram of biomass: 19.7 mg Cu(II), 4.11 mg Ni(II), 7.1 mg Cd(II), 7.7 mg Cr(III), 43 mg Pb(II), and 4.9 mg Zn(II). Most of these metals were recovered from the biomass by treatment with 0.1M HCl with the exception of Cr(III). Because no Cr(VI) binding occurred, none was recovered. Direct and indirect approaches were applied to study the possible mechanisms involved in metal binding by the alfalfa biomass. The direct approach involves investigations of the alfalfa shoot biomass by X-ray absorption spectroscopic analysis (XANES and EXAFS), which were performed at Brookhaven National Laboratory. Results from these studies suggest that Ni(II) binding may occur through coordination with oxygen and some nitrogen ligands, and Cr(III) binding may be occurring through coordination with oxygen ligands. Indirect approaches consist of chemical modification of carboxylate groups which have shown to play an important role in metal binding to the alfalfa biomass. An appreciable decrease in metal binding resulted from acidic methanol esterification of the biomass indicating that carboxyl groups are entailed in the metal binding by the alfalfa biomass. In addition, base hydrolysis of the alfalfa biomass increased the binding of these metals, which further indicates that carboxyl groups play an important role in the binding of Cu(II), Ni(II),Cd(II),Cr(III), Pb(II), and Zn(II) from solution. These studies are important for determining the ligands that may be involved in the binding of metal ions to the alfalfa biomass, thus aiding in the innovative removal and recovery of metal ions from contaminated waters through phyto-filtration.
Keywords: phyto-filtration, alfalfa, medicago sativa, metal ligands, heavy metal binding
There has been an increased concern for public health due to heavy metal contamination in the aqueous environment (Runnells, et al., 1992). Traditional methods for heavy metal removal from contaminated waters involve filtration, flocculation, activated charcoal, and ion exchange resins which are costly and can result in hazardous heavy metal exposure to the workers involved. Although remediation of polluted waters are necessary, in order to prevent further discharge of contaminants into the environment, simpler cost-effective methods need to be developed for industrial use. Currently, a great deal of interest has been given to the use of biological systems for the removal of heavy metals from polluted sites (Baker, et al., 1994; Chamberlain and Miller, 1982; Bewley, et al., 1980; Zhang and Majidi, 1993; Cervantes, et al., 1994; J.R. Lujan, et al., 1994; Gardea-Torresdey, et al., 1990; Gardea-Torresdey, et al., 1996a,1996b).
The use of live biological systems works well with low contaminant levels, but they fall prey to high concentrations of heavy metals (Bender, et al., 1994; Nagendra Rao, et al., 1993). These systems are not as applicable in the harsh conditions of high level industrial waste. Dead or inactivated systems have the advantage of not being affected by high levels of contamination and can also be obtained more cheaply. Several studies have shown that nonliving natural materials are effective for the removal of heavy metals from the environment (Viraraghaven, et al., 1993; de Rome and Gadd, 1991; Ramelow, et al., 1993; Lujan, et al., 1994). Gardea-Torresdey and coworkers demonstrated that carboxyl groups found on the cell walls of dead algal biomass are partially responsible for copper binding (Gardea-Torresdey, et al., 1990). This phenomenon has spurred interest in other natural materials that may contain similar functional groups such as higher plant cells. Jakson and coworkers have studied the binding of cadmium(II) to inactivated cells of Datura innoxia, and other researchers have shown that plant cells are capable of binding heavy metals (Jakson, et al., 1987; Micera and Dessi, 1988; Delhaize, et al., 1989; Scott, 1992; Lujan, et al., 1994). The role of the functional groups involved in the binding of heavy metals is still not well understood. By determining how functional groups are responsible for metal binding by plant cells, low cost technologies can be found to help solve these environmental problems.
Medicago sativa (alfalfa) was identified as a potential biological material that can be used for metal adsorption since it has been found growing in fields irrigated with heavy metal-contaminated water, and was reported to accumulate metal concentrations above the tolerance levels for most plants (El-Kherberawy, et al., 1989; Cajuste, et al., 1991; Baligar, et al., 1993; Rechcigl, et al., 1988). Since alfalfa is inexpensive and high in proteins (which contain many functional groups that could be responsible for heavy metal binding), it was adopted for this study. Previously performed studies have shown that the alfalfa shoot biomass can bind an appreciable amount of copper(II), nickel(II), cadmium(II), chromium(III), lead(II), and zinc(II) ions from aqueous solution (Gardea-Torresdey, et al., 1996b,1996c,1997). Of the seven different alfalfa populations formerly studied, Malone and African exhibited the highest capacity for metal binding. Batch laboratory experiments were performed to determine the pH profiles, time dependency, and capacity for metal binding, as well as the recovery of the metals bound. For most of the metal ions studied, the biomass showed a high affinity for metal binding around pH 5.0 within a short time period. Binding capacity experiments revealed the following amounts of metal ions bound per gram of biomass: 19.7 mg Cu(II), 4.11 mg Ni(II), 7.1 mg Cd(II), 7.7 mg Cr(III), 43 mg Pb(II), and 4.9 mg Zn(II)(Gardea-Torresdey, et al., 1996b,1996c,1997). Most of these metals were recovered from the biomass by treatment with 0.1M HCl, with the exception of Cr(III).
Although much information has been gained on the binding of metal ions to alfalfa biomass, no information is available in relation to the actual metal binding chemical groups. The objective of this study is to identify the possible mechanisms involved in metal binding by the alfalfa biomass. In this report we discuss the characterization of Cr(III) and Ni(II) binding with alfalfa shoot biomass by X-ray absorption spectroscopic analysis (XANES and EXAFS), which was performed at Brookhaven National Laboratory. Also, results from the esterification of functional carboxyl groups and their effects on metal binding are reported. In addition, the effects of alfalfa biomass hydrolysis are reported. These studies are particularly important for determining the ligands that may be involved in the binding of metal ions to the alfalfa biomass, thus aiding in the innovative removal and recovery of metal ions from contaminated waters through phytofiltration.
The Malone population of alfalfa biomass was selected from previous studies for its abundance and metal binding abilities. The plant tissues were collected from controlled agricultural fields at New Mexico State University near Las Cruces, New Mexico. The plants were removed from the soils, washed throughly to remove any debris, and the roots were separated from the shoots (stems and leaves). The samples were then oven dried at 90oC for one week. The dried samples were ground to pass through a 100-mesh screen by using a Wiley Mill.
Esterification of Alfalfa Biomass
Nine grams of Malone alfalfa shoot biomass (oven dried and ground to 100 mesh) were weighed and washed twice with 0.01M hydrochloric acid (HCl) followed by centrifugation for five minutes at 3,000 rpm. This was done to remove any debris and soluble materials. The washings were decanted, dried, and weighed to account for any loss of biomass. The pelleted biomass was next washed with deionized (DI) water three times followed by centrifugation to remove the acidity from the biomass. Again, washings were decanted, dried, and weighed to account for any loss of biomass. The biomass was next suspended in an acidic methanol solution (633 mL of 99.9% methanol and 5.4 mL of concentrated HCl ) which had a final acidic concentration of 0.1M HCl. The solution was continuously stirred and heated to 60 oC for 48 hours. The biomass was then pelleted by centrifugation and the supernatant was decanted, dried, and weighed to account for any loss of biomass. The pellets were then washed with DI water three times using centrifugation as indicated above in order to quench the esterification reaction. The pelleted biomass was then lyophilized for further metal binding experiments.
In addition, controls were maintained from the original biomass and washed with 0.01M HCl and DI as indicated above. One control was reacted with 0.1M HCL, another with DI water, while a third was reacted with pure 99.9% methanol. Each control biomass solution was treated as the first sample and continuously stirred and heated to 60oC for 48 hours. The biomass was then pelleted by centrifugation and the supernatants were decanted as described above. The pellets were then washed with DI water three times using centrifugation as indicated above. All of the control biomass pellets were lyophilized for further metal binding experiments.
Hydrolyzation of Alfalfa Biomass
Nine grams of Malone alfalfa shoot biomass (oven dried and ground to 100 mesh) were weighed and washed twice with 0.01M HCl, followed by centrifugation for five minutes at 3,000 rpm as described above. The biomass was then reacted with 100 mL of 0.1M Sodium Hydroxide (NaOH) for one hour. The biomass was pelleted by centrifugation and the supernatant was decanted as described above. The pelleted biomass was then washed with DI water three times using centrifugation as indicated above, and lyophilized for further metal binding experiments.
Metal Binding Experiment
For each of the hydrolyzed, esterified, and control samples described above, ten milligrams were weighed and placed into clean test tubes. This was performed three times for quality control. Separate metal solutions were made of 0.3mM nickel (from Ni(NO3)2), 0.3mM copper (from CuSO4), 0.3mM cadmium (from Cd(NO3)2.4H2O), 0.3mM chromium (from Cr(NO3)3.9H2O), 0.3mM lead (from Pb(NO3)2), and 0.3mM zinc (from ZnCl2). Each of the metal solutions was adjusted to pH 5.0. Two mLs of 0.3mM metal solution were added to each biomass containing tube and equilibrated for 10 minutes by rocking (final biomass concentration was 5 mg/mL). The biomass was then pelleted by centrifugation for five minutes at 3,000 rpm. The supernatants were kept for analysis and the biomass was again reacted with fresh 0.3 mM metal solution. This process was continued for ten times or until the biomass became saturated and was no longer able to bind more metal from the solution.
X-ray Absorption Spectroscopic Studies
The X-ray absorption spectra were measured at room temperature at beam lines X-18B and X-19A at the National Synchrotron Light Source. Data were collected with Si 111 (X-18B and X-19A) monochromator crystals with slits adjusted to give ~ 1-2 eV resolution. The Ni(II) (H2O)6(NO3)2 standard was measured as an aqueous solution in transmission mode using standard ion detectors. The K2Cr(VI)2O7 and Cr(III)(NO3)3 were measured as solids on tape in transmission mode. Alfalfa biomass immobilized in a silica support matrix as described by Gardea-Torresdey and coworkers was saturated with 1000 ppm of metal solution prior to the analysis (Gardea-Torresdey, et al., 1996b). All of the metal biomass samples were then washed with D.I. water prior to use and run as solid powders in fluorescence mode using a Lytle ionization detector. The absolute energy positions were calibrated with Ni (8333 eV) and Cr (5989 eV) metal foils. The data was analyzed with the MacXFAS EXAFS analysis package using standard methods (Furenlid, et al.,1995). The E0 values were determined from the absorption edge step midpoint. A linear pre-edge background and a least squares cubic spline EXAFS background were extrapolated to normalize the X-ray Absorption Near Edge Spectra (XANES) absorption intensities. Quantitative comparisons between unknown and standards were accomplished with nonlinear fits based on the general Extended X-ray Absorption Fine Structure (EXAFS) equation and verified with theoretical simulations carried out with FEFF 3.11, and ab initio curved-wave, single scattering EXAFS simulation code (Rehr, et al., 1991). Analysis of the EXAFS data for standards was in good agreement with published x-ray crystallographic data.
Each metal studied was analyzed with a Perkin Elmer model 3110 Atomic Absorption Spectrometer with deuterium background subtraction. The methods and conditions followed for each metal analysis were obtained from the Perkin Elmer model 3110 Atomic Absorption Spectrometer manual. Analytical wavelengths used for the various metals were 327.4 nm for copper, 228.8 nm for cadmium, 359.4 nm for chromium, 283.3 nm for lead, 352.5 nm for nickel, and 213.9nm for zinc. Calibration of the instrument was performed within the range of analysis and a correlation coefficient for the calibration curve of 0.98 or greater was obtained. Periodically the instrument's response was checked throughout the analysis with known standards. Samples were read three times and a mean value and relative standard deviation were computed. The difference between the initial control metal concentration and that observed in the supernatant was assumed to be bound by the alfalfa biomass.
RESULTS AND DISCUSSION
Metal Binding Study with Esterified Biomass
Since carboxyl groups have been found to play a role in the binding of heavy metals, acidic methanol was used to chemically block the carboxyl groups by transforming them into methyl esters which should be unable to bind metal ions. A summary of the chemical esterification reaction is shown below.
R-COOH + CH3OH + H+ ® R-COO-CH3 + H2O
In order to determine the extent to which the biomass had been esterified, it was necessary to have non-esterified controls for comparison. These controls consisted of washed biomass reacted with 0.1M HCl, DI water and pure 99.9% methanol, respectively. The methanol and acid controls indicated that the esterification of the biomass was neither due to the acid nor to the methanol alone, but both combined (data not shown). Table 1 shows the mg of metal bound by one gram of alfalfa shoot biomass before and after the esterification reaction. In addition, Table 1 shows the percent decrease in metal binding by the esterified biomass. It can be observed from this table that the addition of acidic methanol to the biomass caused a reduction of 100% of the binding of Cr(III), Pb(II), and Zn(II) by the esterified biomass as compared to the non-esterified (DI) control. Since the acidic methanol treated biomass did not bind any of the Cr(III), Pb(II), and Zn(II) after esterification, carboxyl groups should be the main ligand involved in the binding of these metals. However, since the modified biomass binding was reduced by 93 % for Ni(II), 82% for Cu(II), and 53 % for Cd(II), as compared to the non-esterified (DI) control, other groups may be involved in the binding of these metals. The reduction of the metal binding by the addition of acidic methanol suggests that the carboxyl groups were modified and therefore may play a significant role in the binding of Cu(II), Ni(II),Cd(II),Cr(III), Pb(II), and Zn(II) to the alfalfa biomass.
Metal Binding Study with Hydrolyzed Biomass
The ligands present in the biomass which contain esters and are not active in the binding of metals might be hydrolyzed with 0.1M NaOH, forming new carboxyl groups. A summary of the chemical hydrolyzation reaction is shown below.
R-COO-CH3 + NaOH ® R-COO- + CH3OH
Therefore, by hydrolyzing these esters, we may form new binding sites that were previously unavailable to the metal ions. Table 2 shows the mg of metal bound by one gram of alfalfa shoot biomass before and after the hydrolyzation reaction. In addition, Table 2 shows the percent increase in metal binding by the hydrolyzed biomass. It is observed from Table 2 that the binding was enhanced for the hydrolyzed biomass; 29% for Ni(II), 46 % for Cd(II), 68% for Pb(II), 90% for Zn(II), 111% for Cr(III), and 113% for Cu(II), as compared to the non-hydrolyzed (DI) control. This indicates that the increased metal binding is due to the newly formed carboxyl groups. Thus, the inhibition of metal binding due to blockage of the carboxyl group by acidic methanol addition, and enhancement of metal binding due to ester hydrolyzation, imply that carboxyl groups are responsible for a great portion of the binding of Cu(II), Ni(II), Cd(II), Cr(III), Pb(II), and Zn(II) by the alfalfa biomass.
Metal Binding Studies by X-ray Absorption Spectroscopy
X-ray absorption spectroscopy (XAS) was also used to investigate the possible chemical functional groups responsible for the binding of metal ions to the silica-immobilized alfalfa biomass. XAS is based on the absorption of high-energy monochromatic x-rays by an element in the region of characteristic absorption edge (Lytle, 1988). As ejected photoelectrons from the absorbing atoms scatter and / or make transitions, the effect produces a structure / chemical probe of a few angstroms around the absorbing element. X-ray absorption near-edge structure (XANES) is a probe of the valence and symmetry of the atomic site. As more energetic photoelectrons leave the absorbing atom and are backscattered by the surrounding near-neighbor atoms, it produces a diffraction-like effect called X-ray absorption fine structure (EXAFS). By using EXAFS, one can determine the valance and chemical structure (e.g., coordination numbers, next nearest neighbor information, bond angles, and distances) of elements within a particular chemical environment (Lytle, et al., 1996). Analysis of EXAFS oscillations typically provide accuracies for the coordination number of ±1 and distances of ±0.02Å. XAS experiments were performed for both nickel- and chromium-loaded silica immobilized biomass.
The XANES spectra for the Ni-biomass and NI(II)(H2O)6(NO3)2 have similar edge positions indicating that the nickel binds to the biomass as Ni(II), see Figure 1. The absence of a 1s-4p peak and a weak 1s-3d peak in the XANES spectra for both samples is characteristic of a six-coordinate octahedral environment for the nickel. The isolated k3 EXAFS data and the Fourier Transform (FT) -EXAFS data are shown in Figures 2 and 3, respectively. Good fits are obtained using 6 nitrogens at 2.08Å, 5 oxygens at 2.05Å, or a mixture of nitrogen and oxygen atoms. Fits using only sulfur or a mixture of sulfur and oxygen or nitrogen all gave unreasonable results. From these XAFS studies we can say that the nickel is six-coordinate with either nitrogen or oxygen ligands. The lack of multiple scattering peaks at 2.93Å and 4.24Å, see Figure 3, associated with imidazole carbon and nitrogen atoms, suggests that if nickel coordinates via nitrogen atoms they are not histidines. Coordination of the nickel by sulfur is not evident, since one would expect long Ni-S bonds of ~2.2 Å. The small Debye-Waller factors fail to indicate a multitude of different nickel binding sites of different geometries.
The normalized XANES spectra for Cr-biomass, Cr(III)(NO3)3(H2O)6, Cr(III)2S3, and K2Cr(VI)2O7 are shown in Figure 4. The edge position for the Cr-biomass and Cr(III)(NO3)3(H2O)6 and Cr(III)2S3 are - 5eV relative to the Cr(VI) standard and the absence of a strong pre-edge transition, characteristic of Cr(VI), indicate that the chromium binds to the biomass as Cr(III). Comparison of the Cr-biomass with the six coordinate Cr(III) standards containing oxygen or sulfur ligands strongly suggests that the Cr is octahedral with oxygen axial ligands. The isolated EXAFS and FT-EXAFS data are shown in Figures 5 and 6, respectively. The EXAFS oscillations and transforms for the Cr-biomass closely resemble the Cr(III)(NO3)3(H2O)6 standard. The best single shell fit for the isolated first shell EXAFS oscillations was for 5 oxygens at 2.00Å; both nitrogen and sulfur atoms gave worse fits. The two shell fits using 4 oxygen and 1 nitrogen atoms afforded better fits, but had an unreasonably short Cr-N distance of 1.92Å. Both the XANES and EXAFS data indicate that the Cr-biomass is six coordinate with oxygen ligands at 2.00Å. The small Debye-Waller factors, as found for the Ni-biomass, also indicate no multitude of Cr binding sites with different geometries.
Both the indirect and direct methods employed have provided clues to the mechanism of metal binding to alfalfa shoot tissues. Chemical modification of the alfalfa biomass by esterification of available carboxyl ligands with acidic methanol has shown a dramatic reduction in the biomass metal interaction for Cu(II) ,Ni(II),Cd(II),Cr(III), Pb(II), and Zn(II). In addition, hydrolysis of the alfalfa biomass can appreciably increase the binding of these metals, which further indicates that carboxyl groups play an important role in the binding of Cu(II) ,Ni(II),Cd(II),Cr(III), Pb(II), and Zn(II) from solution. Also, XANES and EXAFS results from these studies suggest that for both Ni and Cr biomass complexes, the metal binding may occur through coordination with oxygen or nitrogen ligands. Although amino, sulfhydryl, and hydroxyl groups must be considered, these results have shown that carboxyl groups may be responsible for a large portion of the metal biomass interaction. Currently we are performing experiments to determine XANES and EXAFS of other metals bound to the alfalfa biomass. By determining the actual chemical functional groups responsible for the binding of heavy metals, we will be able to assist in producing innovative technologies for the removal and recovery of heavy metals from contaminated waters through phytofiltration.
The authors acknowledge financial support from the National Institutes of Health (NIH), (grant # GM 08012-25) and financial support from the University of Texas at El Paso's Center for Environmental Resource Management (CERM) through funding from the Office of Exploratory Research of the U.S. Environmental Protection Agency (cooperative agreement CR-819849-01-4). We also acknowledge the HBCU/MI Environmental Technology Consortium which is funded by the Department of Energy. The work performed at Brokhaven National Laboratories was supported by the Department of Energy, Chemical Sciences Division (Contract DE-AC02-76CH00016).
Baker, A.J.M., R.D.Reeves, and A.S.M.Hajar, 1994. Heavy Metal Accumulation and Tolerance in British Populatins of the Metallophyte Thalspi caerulescens (Brassicaceae), J. & C. Pres, New Phytol., 127, pp.61-68.
Baligar,V. C., T.A.Campbell and R.J. Wright, 1993. Differential Responses of Alfalfa Clones to Aluminum-Toxic Acid Soil, Plant Nutrition, 16, pp. 219-233.
Bender, J., S. Rodriguez-Eaton, U.M. Ekanemesang and P. Phillips, 1994. Characterization of the Metal Binding Bioflocculants Produced by the Cyanobacterial Component of Mixed Microbial Mats, App. Environ. Microbiol., 60, pp. 2311-2321.
Bewley, R.J., 1980. Effects of Heavy Metal Pollution on Oak Leaf Microorganism, App. Environ. Microbiol., 40, pp.1053-1059.
Cajuste, L.J., R. Carrillo, G.E. Cota and R.J. Laird, 1991. The Distribution of Metals from Wastewater in the Mexican Valley of Mezquital, Water, Air and Soil Pollution., 57, pp.763-771.
Cervantes,C. and F. Gutierrez-Corona, 1994. Copper Resistance Mechanisms in Bacteria and Fungi, FEMS Microbiol. Rev., 14, pp.121-138.
Chamberlain,W.F. and J.A. Miller,1982. Barium in Forage Plants and in the Manure of Cattle Treated with Barium Boluses, J. Agric. Food Chem., 30, pp.463-465.
Delhaize, E., P.J. Jackson, L.D. Lujan and N.J. Robinson, 1989. Poly (g- glutamylcysteinyl) glicine Synthesis in Datura innoxia and Binding with Cadmium, Plant Physio., 89, pp.700-706.
El-Kherbawy, M., J.S. Angle, A. Heggo and R.L. Chaney, 1989. Siol pH Rhizobia, and Vesicular-arbuscular Mycorrhizae Innoculation Effects on Growth and Heavy Metal Uptake of Alfalfa (Medicago sativa), Biol. Fertil. Soils, 8, pp.61-73.
Furenlid, L.R., M.W. Renner and E. Fujita, 1995. XAS Studies of Ni(I), Ni(II), and Ni(III) Complexes, Physica B., 208 & 209, pp.739-742.
Gardea-Torresdey, J.L., M.K. Becker-Hapak, J.M. Hosea, and Dennis W. Darnall, 1990. Effect of Chemical Modification of Algal Carboxyl Groups on Metal Ion Binding, Environ. Sci. Technol., 24, pp.1372-1378.
Gardea-Torresdey, J.L., I. Cano-Aguilera, R. Web, K.J. Tiemann and F. Gutierrez-Corona, 1996a. Copper Adsorption by Inactivated Cells of Mucor rouxii:Effect of Esterification of Carboxyl Groups, J. of Haz. Mat., 48, pp.171-180.
Gardea-Torresdey, J.L., K.J. Tiemann, J.H. Gonzalez, J.A. Henning and M. S.Towsend, 1996b. Ability of Silica-Immobilized Medicago Sativa (Alfalfa) to Remove Copper Ions from Solution, J. of Haz. Mat., 48, pp.181-190.
Gardea-Torresdey, J.L., K.J. Tiemann, J.H. Gonzalez, J.A. Henning and M. S.Towsend, 1996c. Removal of Nickel Ions from Aqueous Solution by Biomass and Silica-Immobilized Biomass of Medicago Sativa (Alfalfa), J. of Haz. Mat., 49 pp.205-216.
Gardea-Torresdey, J.L., K.J. Tiemann, J.H. Gonzalez, O. Rodriguez and G. Gamez, 1997. Phytofiltration of Hazardous Cadmium, Chromium, Lead, and Zinc Ions by Biomass of Medicago Sativa (Alfalfa), J. of Haz. Mat., (in press).
Jakson, P.J., C.J. Unkefer, J.A. Doolen, K. Watt, and N.J. Robinson, 1987. Poly (g-glutamylcysteinyl) glicine: Its Role in Cadmium Resistance in Plant Cells, Proc. Natl. Aca Sci., 84, pp.6619-6623.
Lujan, J.R., D.W. Darnall, P.C. Stark, G.D. Rayson and J.L. Gardea-Torresdey, 1994. Metal Ion Binding by Algae and Higher Plant Tissues: A Phenomenological Study of Solution pH Dependence, Solvent Extr. Ion Exch., 12, pp.803-816.
Lytle, C.M., F.W. Lytle, and B.N. Smith, 1996. Use of XAS to Determine the Chemical Speciation of Bioaccumulated Manganese in Potamogeton pectinatus, J. Environ Qual., 25, pp.311-316.
Lytle, F.W., 1988. Applications of Synchrotrn Radiation, Gordon and Breach, Newark, NJ., pp.87-102.
Micera, G. and A. Dessi,1988. Chromium Adsorption by Plant Roots and Formation of Long-lived Cr(VI) Species: An Ecological Hazard?, J. Inorg. Biochem., 34, pp.157-166.
Nagendra Rao, C.R., L. Iyengar and C. Venkobachar, 1993. Sorption of Copper II from Aqueous Phase by Waste Biomass, J. Environ. Eng., 119, pp.369-377.
Ramelow, G.J., L. Liu, C. Himel, D. Fralick, Y. Zhao and C. Tong, 1993. The Analysis of Dissolved Metals in Natural Waters after Preconcentration on Biosorbents of Immobilized Lichens and Seaweed Biomass in Silica, Intern J. Anal. Chem., 53, pp.219-232.
Rechcigl, J.E., R.B. Reneau and L.W. Zelazney, 1988. Soil Solution Al as a Measure of Al Toxicity to Alfalfa in Acid Soils, Soil Sci. Plant Anal., 19, pp.989-1001.
Rehr, J.J., M. Balci, K. Pramod, P. Koch, J. Lex and O. Ermer, 1991. Theoretical X-ray Absorption Fine Structure Standards, J. Am. Chem. Soc.,113, pp.5135-5149.
Rome, L. and G.M. Gadd, 1991. Use of Pelleted and Immobilized Yeast and Fungal Biomass for Heavy Metal and Radionuclide, J. Industrial Microbiol., 7, pp.97-104.
Runnells, D.D., T.A. Shepherd and E.E. Angino, 1992. Metals in Water: Determining Natural Background Concentrations in Mineralized Areas, Environ. Sci. Technol., 26, pp.2316-2323.
Scott, C.D., 1992. Removal of Dissolved Metals by Plant Tissue, Biotechnol. and Bioeng., 39, pp.1064-1068.
Viraraghaven, T., R. Saskatchewan and M.M. Dronamraju, 1993. Removal of Copper, Nickel and Zinc, from Wastewaters by Adsorption Using Peat, J. Environ. Sci. Health, A28, pp.1261-1269.
Zhang W. and V. Majidi, 1993. Study of Influences on the Binding of Metals to Stichococcus bacillaris with 113Cd NMR, Applied Spectroscopy, 47, pp.2151-2158.
Table 1. Capacity for metal binding by chemically modified alfalfa shoots using acidic methanol (before and after esterification).
mg metal/ g Alfalfa biomass
|BEFORE ESTERIFICATION||AFTER ESTERIFICATION||% DECREASE IN BINDING|
Table 2. Capacity for metal binding by chemically modified alfalfa shoots using sodium hydroxide (before and after hydrolysis)
mg metal/ g Alfalfa biomass
|BEFORE HYDROLYSIS||AFTER HYDROLYSIS||% INCREASE IN BINDING|
Figure 1. X-ray absorption near-edge spectra for Nickel bound biomass (solid) and
Ni(H2O)6(NO3)2 (solution) at room temperature.
Figure 2. Isolated k3-weighted EXAFS oscillation for Nickel bound biomass (solid) and
Ni(H2O)6(NO3)2 (solution) at room temperature.
Figure 3. Fourier transform magnitudes of the k3-weighted EXAFS oscillation for Nickel
bound biomass (solid) and Ni(H2O)6(NO3)2 (solution) at room temperature.
Figure 4. X-ray absorption near-edge spectra for Chromium bound biomass (solid),
K2Cr2O7 , Cr(III)2S3 and Cr(III)(NO3)3 (solids) at room temperature.
Figure 5. Isolated k3-weighted EXAFS oscillation for Chromium bound biomass (solid),
K2Cr2O7, Cr(III)2S3 and Cr(III)(NO3)3 (solids) at room temperature.
Figure 6. Fourier transform magnitudes of the k3-weighted EXAFS oscillation for
Chromium bound biomass (solid), D2Cr2O7, Cr(III)2S3, and Cr(III)(NO3)3 (solids) at room