**1. Introduction**

140 Biomaterials – Physics and Chemistry

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Removal of toxic heavy metal ions from contaminated water is required to provide safe drinking water. This can be effected either within the waste stream at contaminate source or at point of use. Incorporation of remediation technologies at either location requires the removal of pollutants at parts per million and parts per billion concentrations from water containing more benign metal ions (e.g., Ca2+, Mg2+, and Na+) at concentrations three to six orders of magnitude higher. Materials derived from plants or microorganisms (e.g., algae and fungi) have been shown to enable the reduction of trace concentrations of heavy metal ions to below regulatory limits (Davis, et al., 2003).

Such nonliving biomaterials have been reported to have exhibit high capacity, rapid binding, and selectivity towards heavy metals (Drake and Rayson, 1996). It is postulated that functional groups native to the lipids, carbohydrates, and proteins found in the cell walls of the biomaterial are responsible for uptake (biosorption) of metal ions (Gardea-Torresdey, et al., 1999; 2001; Drake and Rayson, 1996; Drake, et al., 1997; Kelley, et al., 1999). For biomaterials to become a commercially viable method of metal remediation and recovery these functional groups must be identified and their contribution to overall metal binding capacity quantified. Knowledge of such informaiton would allow either simple chemical alteration to the biomaterial, allowing for targeting of specific metals, or an enhancement of biomaterial metal binding.

Significant progress has been made to identify the chemical functionalities involved in the biosorption of numerous metal ions by a variety of plant and algal tissues (Gardea-Torresdey, et al., 1999; 2001; Riddle, et al., 2002; Fourest and Volesky, 1996; Drake, et al., 1997;Jackson, et al., 1993). Several techniques have been reported to probe local chemical environments of biosorbed metal ions. These have included X-ray absorption (Gardea-Torresdey, et al., 1999; Riddle, et al., 2002), lanthanide luminescence (Drake, et al., 1997; Serna, et al., 2010), and metal NMR (Xia and Rayson, 1996; 2002; Kelley, et al., 1999; Majidi, et al., 1990) spectroscopy. Analysis of total metal ion binding isotherm data modeling (Volesky, 2000) has also been described. Efforts to address the chemical heterogeneity of those biosorbed materials have also employed regularized regression analysis of isotherm data (Lin, et al., 1996). Additionally, these chemical intensities have been studied through selective removal of binding moieties by their reactive modification (Drake, et al., 1996).

Comparative Metal Ion Binding to Native

ensure complete sulfate removal.

**2.3 Biomaterial columns** 

environment (~pH 6.2).

other) metal species in the column effluent.

and 75.3% for the native and modified materials, respectively.

observed and their respective emission wavelengths.

and Chemically Modified *Datura innoxia* Immobilized Biomaterials 143

further raised to 7.0 by incremental addition of the Na2SiO3 solution. A gelatinous polymer formed at pH 7.0. The solution was stirred an additional 30 minutes, covered and stored at 4°C overnight. The resulting aqueous layer was removed. Excess sulfate ions were removed from the gel by successive washings with distilled deionized water until the aqueous phase failed a sulfate test using a few drops of a 1.0% barium solution (as the nitrate salt). When no precipitate was visible, one final wash with the distilled deionized water was performed to

Remaining polymer was transferred to ceramic evaporating dishes and baked at approximately 100 C until completely dry. The immobilized *D. innoxia* biomaterial was then ground and sieved. The 40-60-mesh (423-635 μm) particle size fraction was collected. Percent compositions of cell wall biomaterial were determined gravimetrically to be 64.6%

The columns used have been described elsewhere (Williams and Rayson, 2003) and were constructed in-house from Plexiglas™ tubing (2.5 cm in length and 3 mm i.d.). Teflon™ tubing (0.8-mm i.d.) was used for all column connections. Interface of the column to the ICP-OES was accomplished by connecting the column outlet directly to the inlet of the crossflow type nebulizer using the minimum length of Teflon™ tubing (15 cm). Column effluent was monitored for each of 27 different metals simultaneously. Table 1 list the elements

Element Wavelength/nm Element Wavelength/nm Element Wavelength/nm Na 589.00 Mg 279.00 Al 396.15 Ca 317.90 Cr 267.70 Mn 257.60 Fe 259.90 Ni 231.60 Cu 324.70 Zn 213.80 Cd 228.80 Ag 328.00 Sn 189.90 Pb 220.30 Ba 493.40 Sr 421.50 U 409.00 Y 371.00 V 242.40 Mo 202.00 Co 228.60 Si 251.60 As 193.60 Se 196.00 Tb 350.97 Eu 381.97 Th 283.73 Table 1. Elements and the corresponding emission wavelength used during monitoring of

column effluents (elements of interests in this study indicated by boldface print).

Each column was packed with approximately 125 mg of the immobilized *D. innoxia* material and flow tested using distilled deionized water. Once packed and tested for leaks, each column was exposed to 20 mL of 1.0-M HCl using a peristaltic pump (Model Rabbit, Rainin) (1.0 mL/min for 20 min) and the effluent monitored for metals released from the biomaterial. Following the acid rinse, the columns were then exposed to 5 mL of distilled deionized water (1.0 mL/min for five minutes) to reestablish an ambient pH influent

These studies involved, initially, the exposure of a small column (3.0 mm i.d., 10.0 mm in length) to an equimolar mixture of metal ions, specifically, Cd2+, Zn2+, and Ni2+, and exposure to solutions of each metal sequentially while continuously monitoring these (and

It has been shown that carboxyl groups present in the cell walls of nonliving biomaterials contribute to metal-ion binding (Gardea-Torresdey, et al., 1999; 2001; Riddle, et al., 1997; Kelley, et al., 1999). Our group has used a variety of methods(Lin, et al., 2002; Drake, et al., 1996; Xia and Rayson, 1995; Drake, et al., 1996; 1997) to characterize the binding groups present in the cell walls of *Datura innoxia.* This plant is a member of the *Solanaceae* plant family and native to Mexico and the southwestern United States. To minimize variability of cell types investigated, the cell-wall fragments used are cultured anther cells of the plant. This plant was selected for study because it is a heavy metal resistant perennial that is both tolerant of arid climates and resistant to herbivory (Drake et al., 1996).

Our group has concentrated primarily on nonviable biomaterials, specifically cell wall fragments from the cultured anther cells of *Datura innoxia.* The present study used frontal affinity chromatography with inductively coupled plasma optical emission spectroscopy (ICP-OES) detection for simultaneous monitoring both uptake and release of metal ions to both a chemically modified and native *D. innoxia* biomaterial (Williams and Rayson, 2003). The objective of the present study was to further investigate such sites through sequential exposure and subsequent stripping of three similar metal ions (Cd2+, Ni2+, and Zn2+) to both a modified and the native biosorbents, thus to study the role of carboxylate furface functionalities on passive metal ion binding of this material.

It has been demonstrated (Drake, et al., 1996) that carboxylate-containing binding sites can be removed through the formation of the corresponding methyl esters by reaction with acidic methanol for 72 hours (Drake, et al., 1996). Undertaking a similar series of experiments with such a chemically modified sorbent enables the investigation of alternate binding sites.

#### **2. Materials and methods**

#### **2.1 Esterification of biomaterial**

The cultured anther cells from *D. innoxia* were washed and prepared as described elsewhere (Drake, et al,, 1996; 1997). Only cell fragment aggregates with a mesh size greater than 200 (< 127 μm) were used for esterification. Following a method described elsewhere (Drake, et al., 1996), 10.0 grams of the biomaterial were suspended in 0.1 M HCl in methanol. The slurry was continuously heated at 60C and stirred for 72 hours. The biomaterial was then recovered through vacuum filtration, rinsed three times with 16.0-M water (Barnstead,Millipore Ultrapure), freeze-dried, and set aside for later immobilization.

#### **2.2 Immobilization of biomaterial**

In their native state, biomaterials have poor mechanical strength, low density, and a small particle size that can cause column clogging (Stark and Rayson, 2000). These characteristics can yield poor candidates for column-based water treatment applications. For this study native and modified *D. innoxia* biomaterials were each immobilized in a polysilicate matrix. The 40-60-mesh size fraction of the ground, and sieved immobilized biosorbents was then packed into columns. The process for immobilization has been described in detail elsewhere (Stark and Rayson, 2000).

Briefly, a suspension of 20 grams of the 100-200 mesh fraction of the washed biomaterial was generated with 300 mL of 5% v/v sulfuric acid adjusted to pH 2.0 by addition of a 6% (w/v) solution of Na2SiO35H2O. This suspension was stirred for 1 hour and the pH of the solution further raised to 7.0 by incremental addition of the Na2SiO3 solution. A gelatinous polymer formed at pH 7.0. The solution was stirred an additional 30 minutes, covered and stored at 4°C overnight. The resulting aqueous layer was removed. Excess sulfate ions were removed from the gel by successive washings with distilled deionized water until the aqueous phase failed a sulfate test using a few drops of a 1.0% barium solution (as the nitrate salt). When no precipitate was visible, one final wash with the distilled deionized water was performed to ensure complete sulfate removal.

Remaining polymer was transferred to ceramic evaporating dishes and baked at approximately 100 C until completely dry. The immobilized *D. innoxia* biomaterial was then ground and sieved. The 40-60-mesh (423-635 μm) particle size fraction was collected. Percent compositions of cell wall biomaterial were determined gravimetrically to be 64.6% and 75.3% for the native and modified materials, respectively.
