Europium Uptake and Partitioning in Oat (Avena sativa) Roots as

The uptake of Eu3+ by elongating oat roots was studied by fluorescence ... Distribution of 152Eu and 154Eu in the 'alluvial soil–rhizosphere–plant...
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Environ. Sci. Technol. 2003, 37, 5247-5253

Europium Uptake and Partitioning in Oat (Avena sativa) Roots as Studied by Laser-Induced Fluorescence Spectroscopy and Confocal Microscopy Profiling Technique ROBERT J. FELLOWS, ZHEMING WANG,* AND CALVIN C. AINSWORTH Pacific Northwest National Laboratory, P.O. Box 999, MS K8-96, Richland, Washington 99352

The uptake of Eu3+ by elongating oat roots was studied by fluorescence spectroscopy, fluorescence lifetime measurement, and a laser excitation time-resolved confocal fluorescence profiling technique. The results of this work indicated that initial uptake of Eu3+ was highest within the undifferentiated cells of the root tip just behind the root cap, a region of maximal cell growth and differentiation and with incomplete formation of the Casparian strip around the central vascular cylinder. Distribution of assimilated Eu3+ within the root’s differentiation and elongation zone was nonuniform. Higher concentrations of Eu3+ were observed within the vascular cylinder, specifically in the phloem and developing xylem parenchyma. Elevated levels of the metal were also observed in the root hairs of the mature root zone. Fluorescence spectroscopic characteristics of the assimilated Eu3+ suggested that the Eu3+ exists as inner-sphere mononuclear complexes inside the root. This work also demonstrated the effectiveness of a time-resolved Eu3+ fluorescence spectroscopy and confocal fluorescence profiling techniques for the in vivo, realtime study of metal [Eu3+] accumulation by a functioning intact plant root. This approach can prove valuable for basic and applied studies in plant nutrition and environmental uptake of actinide radionuclides.

Introduction Recently phytoremediation, or more specifically phytoextraction, has been viewed as a significant economical decontamination method for large volumes of soils and subsurface sediments diffusely contaminated with either radioactive or nonradioactive inorganic contaminants. However fundamental processes such as those factors which govern the rate of metal ion uptake, determine cellular specificity, modify uptake and transport mechanisms, and control cellular and tissue partitioning of heavy metals within the roots and shoots of higher plant roots remain to be identified (1-5). Typical plant metal-uptake experiments involve multiple processes such as plant harvesting, drying, fixation, grinding, and extraction. While these techniques may quantitatively determine total metal ion uptake and/or the general metal location within the plant or root, they often lack cellular and tissue specificity within the root. * Corresponding author phone: (509)376-6119; fax: (509)376-3650; e-mail: [email protected]. 10.1021/es0343609 CCC: $25.00 Published on Web 10/17/2003

 2003 American Chemical Society

The majority of studies on localization of heavy metals within specific tissues have been performed in vitro and have included autoradiography of radioactive isotopes (6) as well as electron probe X-ray microanalysis of individual ions (79). While the latter technique provides enhanced resolution of water-soluble ions within specific cell types of the root, both require that the tissue be fixed or stabilized either under the photographic emulsion or within the scanning electron microscope (SEM) thus preventing real-time observations in living tissues. Organic dyes, dependent on free ions such as Ca2+, Mg+, and Zn2+ for their fluorescent properties, are available for imaging these ions within living tissues (10). However these can be difficult to introduce into cells and may alter the physiology of the tissue. The application of green fluorescent protein (GFP) and other fluoresceins of biological origin have permitted the observation of macromolecular transport within living tissues and cells such as sieve elements and pollen tubes (10-12). Modifications of GFP for immunofluorescence or as indicators for pH and calcium ions (13) have also been reported. However, while these large molecules function as indicators of the free ion movement and location, they will not indicate the chemical state(s) of the native ion within the tissue or cells and further, because of their size, may alter the physiological processes. It would therefore be desirable if there were metal ions whose spectroscopic properties varied with the ion’s chemical state and were capable of being accumulated and transported by living cells. One such ion is trivalent europium, Eu(III). Europium(III) absorbs in the UV and near-UV and exhibits sharp fluorescence emission spectra in the visible range with fluorescence lifetimes that range between tens of microseconds to milliseconds (14-16). The fluorescence intensity, relative intensity among the fluorescence bands, and fluorescence lifetime are all sensitive to its chemical environment permitting precise analysis of the ion’s interaction with its immediate surrounding (16, 17). With the use of pulsed lasers and time-resolved detection techniques the long fluorescence lifetime allows Eu(III) fluorescence measurements even in sample matrices containing strongly fluorescent organic or biological fluorophores whose fluorescence lifetimes (nanoseconds) (18-20) are >103-fold shorter than that of Eu(III). Europium(III) is often used as an analogue to Ca(II) and Mg(II), two of the most abundant ions found in living systems (16, 17). These three ions have similar ionic radii, large coordination numbers, a preference to bind to oxygen or nitrogen donor ligands such as amino and carboxylic acids, and generally little or no metal-centered preference in coordination geometry. In biological systems Eu(III) substitution for Ca/Mg does not affect normal physiological functions (16, 17). Europium(III) is commonly used as a representative of the trivalent lanthanides as well as an analogue of the more toxic actinides such as Cm(III) and Am(III) (15). Although not a required plant nutrient, Eu(III) is capable of root uptake and has been shown to be incorporated into the above-ground foliage of plants growing in media containing the element (21-23). Potential phytotoxic effects of Eu(III) are unknown, while some experimental results indicated that application of rare earth elements at trace level increase the yield of cereals, sugar beets, and other plants (24). Here we report on the uptake of Eu3+ by oat (Avena sativa), its distribution among different root tissues, and its chemical state inside the oat root determined through the use of fluorescence spectroscopy, fluorescence lifetime measureVOL. 37, NO. 22, 2003 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 1. A picture of a typical oat plant used in the Eu(III) uptake experiments and diagrams of the sample stage used for microscopic observation of a single root from an intact, transpiring plant exposed to aqueous Eu(III).

TABLE 1. Composition of Oat Root Growth Media macronutrients

molarity

micronutrients

molarity

KNO3 Ca(NO3)2‚4H2O MgSO4‚7H2O NaH2PO4 HCl iron as Fe-EDTA

6 mM 4 mM 2 mM 1 mM 10 µM 10 µM

H3BO3 MnSO4‚H2O ZnSO4‚7H2O CuSO4‚5H2O Na2MoO4‚2H2O

3 µM 0.5 µM 0.4 µM 0.2 mM 0.05 µM

ment, and a laser excitation time-resolved confocal fluorescence profiling technique. The latter was found to be an in-situ, nondestructive, and highly sensitive technique for long-lived fluorescent heavy metal ions.

Materials and Methods Plant Material. Oat plants (Avena sativa cv. Coker) were grown under sterile conditions. Seeds were first dissected from the bracts, surfaced-sterilized in 0.5% (v/v) sodium hypochlorite for 10 min, and washed 3 times with sterile distilled water. They were then aseptically transferred to onehalf strength tryptic soy broth agar plates for 3 days as a check for sterility. Seeds exhibiting no contamination were transferred to a Petri dish containing a sterile filter paper disk moistened with sterile distilled water. After the germinated seedling had exceeded a total length of 6 cm (4-5 days postinhibition) it was transferred in a laminar flow hood to a sterile 5-mL syringe barrel from which the needle end had been removed, and the resulting opening lightly plugged with a small piece of sterile nonabsorbent cotton. The seedling was placed such that the developing root extended 1-2 cm beyond the cotton plug. Two to 3 cm of sterile acid-washed sand previously treated with Life-Guard Waterproofing Sealant (Life-Guard Waterproofing Products, Inc., Anaheim, CA) to prevent moisture wicking was then lightly packed around the seedling for support. The seedling-syringe assembly was in turn sealed with silicone caulk into the screw cap of a sterilized 600-mL glass jar (Figure 1A) containing 500 mL of 0.22 µm filtered one-tenth strength-Fe Hoagland’s solution (pH 5.5) (Table 1). The nutrient solution was constantly aerated with 0.22 µm-filtered air. The high humidity and light spray from the aeration prevented the root from drying out before it grew down into the liquid. The plants were maintained in a growth chamber at a light intensity of 400 µE m-2 s-1 at mid-canopy, a 12/12 h day/night photoperiod, 50% relative humidity, and a 25 °C/ 20 °C day/night temperature. Filter-sterilized iron-EDTA (0.75 mM) was added to the nutrient solutions after 21 days or earlier if Fe-deficiency symptoms became evident. All chemicals used in the experiments were reagent grade from either Aldrich or Sigma (St. Louis, MO). 5248

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Eu(III) Uptake Experiments. Two types of Eu uptake experiments were performed. In the first, 28-day-old seedlings (4 leaf stage) plants were transferred to 0.45 µm filtered nutrient solutions containing either 1 or 10 mM Eu(III)Cl3‚ 7H2O and maintained under the growth conditions described above. In the second experiment, individual 21- or 28-dayold plants were placed directly into a stage-mounted splitcell cuvette constructed of polyetherketone with quartz windows and liquid inlet and outlet to permit exchange of the nutrient solution with minimum disturbance of the plant root under observation (Figure 1B). A single root was placed into one of the cells and the remaining roots and plant placed in the second, larger root reservoir. The cell containing the single root was filled with 0.22 µm filter-sterilized-Fe nutrient solution containing 10 mM Eu(III), while the remaining roots were maintained in the control-Fe nutrient solution. The plant shoot was illuminated (∼200 µE m-2 s-1) between spectroscopy measurements made on the exposed root at 0-, 1-, 4-, 8-, 24-, 32-, 48-, 56-, 72-, 80-, 96-h, and 120-h intervals as described below. Spectroscopy Measurements. Fluorescence excitation and emission spectra of excised roots and leaves of exposed plants were taken on a SPEX Fluorolog II fluorimeter equipped with two double monochromators (SPEX1681), a 450 W xenon lamp, and a cooled Hamamatsu R928 photomultiplier tube detector. In vivo Eu(III) fluorescence lifetime and fluorescence intensity measurements of its distribution inside the intact root and free-hand root cross sections (10-30 µ thick) were measured using a custom-built laser confocal fluorescence microscope system. The system consists of a pulsed Nd:YAG laser (Spectra-Physics Quanta-Ray Pro-190) pumped MOPO laser (Spectra-Physics MOPO-730), a Nikon inverted optical microscope (Eclipse TE-300), a thermoelectrically cooled Hamamatsu R928 photomultiplier, and a Tektronics TDS 754A digital oscilloscope. The 798 nm output of the MOPO laser was frequency doubled using a KDP crystal to generate 394 nm pulsed laser light with pulse width of ca. 10 ns at 20 Hz. The 394 nm light was expanded and spatially filtered using a set of fused silica lenses and pinhole. The collimated laser beam was then directed into the side port of the inverted microscope with a set of mirrors and a long-pass dichroic filter and focused on the plant root by microscope objective lenses with magnification between 4× to 60×. The emitted fluorescence emission from the plant root was collected by the same objective lens and detected by the photomultiplier tube after passing through the dichroic filter, a focusing lens, and a 17.5 µ pinhole and recorded with the oscilloscope. Fluorescence decay curve was averaged with at least 200 laser pulses. The fluorescence intensity of Eu(III) was determined by integration of the fluorescence decay curve

in the time range of 0.14-1.32 ms, after correction of fluorescence baseline. Fluorescence lifetime measurement of the short-lived emission from the oat root and the growth media was performed using a custom-built time-correlated singlephoton-counting apparatus described previously (25, 26). Laser excitation was provided with the frequency-doubled output (λ ) 300 nm) from a Coherent 702 dye laser operating at 600 nm which was pumped with a Mode-locked Coherent Antares Nd:YAG laser. The pulse duration was approximately 100 ps, and the time duration between pulses was adjusted using a Coherent 7220 Cavity Dumper to allow complete fluorescence decay before the start of the next pulse. Fluorescence decay curves for oat root and growth medium were recorded at emission wavelengths of 440 and 460 nm, respectively, with a maximum reading of at least 104 counts/ channel. Data analysis was performed using commercial software from Wavemetrix, IGOR. Fluorescence decay curves were fit as a sum of exponential functions of the form

I)

∑A exp(-k t) i

i

(1)

where the fluorescence intensity I is contributed by one or more time-dependent components with initial intensities Ai and decay constants ki (corresponding to lifetimes τi where τ ) k-1). Taking advantage of the nearly 5 orders of magnitude longer fluorescence lifetime of Eu(III) as compared to that of the root itself, fluorescence measurement of root assimilated Eu(III) was made by introduction of a small timedelay (a few µs) after the laser pulse from the nanosecond MOPO-730 laser operating at 394 nm. During the delay period, the fluorescence of the root itself was decayed away, leaving only the long-lived Eu(III) fluorescence. Such time-resolved measurement coupled with the use of the confocal optical microscope allowed fluorescence measurement of root assimilated Eu(III) with high spatial resolution of up to a few micrometers, enough to resolve various tissues of the oat root. At the time of this study a time-resolved laser scanning confocal optical microscope was not yet available. In this work, the excitation laser was not scanned on the confocal microscope system. Instead, the laser beam was focused with a stationary 60× water-immersion objective lens on the microscope, and the sample which was mounted on an automated translation stage was stepped across the focal point of the laser beam. At each point the fluorescence decay at 616 nm was recorded at a constant exposure time. The fluorescence intensity was calculated by integrating the area under the fluorescence decay curve excluding the short time range which includes the intensity contributions from the flurescence emission from the oat root as well as scattered laser light. Since the fluorescence lifetime of the root assimilated Eu(III) was nearly constant at different locations along the oat root, it is reasonable to assume the same speciation was maintained. Therefore, the amount of assimilated Eu(III) was proportional to the measured fluorescence intensity. White light images of the intact root and cross sections were recorded with a Nikon camera attached to the front port of the microscope in transmission mode and halogen lamp illumination. At the conclusion of the exposure experiments, some of the hand-sectioned tissues were stained with 0.05% (w/v) Toluidine Blue for 3 min and rinsed in distilled water to enhance photographic contrast.

Results and Discussion Plant Growth and Eu(III) Uptake. Under hydroponic conditions (Figure 1A) the oat plants typically grew into a 2-4 leaf plant with shoot length between 10 and 15 cm, and the length of the main root was about 10 cm with mature

FIGURE 2. Steady-state fluorescence emission spectra of (a) excised oat root; (b) the oat plant growth medium; (c) 0.01 M EuCl3 aqueous solution. λex ) 394 nm. The asterisks stands for water Raman band. Note that a direct comparison between the relative intensity of the oat roots and those of the growth media or the aqueous solution of EuCl3 is invalid due to the ill-defined excitation volume of the roots. root diameter of 500 µm-1 mm after 28 days postgermination. The growth media appeared clear throughout the experiment. Microscopically, the plant roots were translucent with no discernible mineral precipitation with or without the presence of Eu(III) chloride (0.001-0.01 M). The roots of both terrestrial and aquatic plants have an intimate association with the microbial inhabitants in their rhizosphere, the area immediately surrounding the root surface (27). Interactions by both plants and microbes may include ligand secretions, e.g. siderophores and phytosiderophores capable of mobilizing nutrient metals such as Fe, Zn, Mg, and Cu (28) as well as organic acids and protons which alter the pH and oxidation state of the microenvironment immediately surrounding the root. Sterile culture of the plants was performed to avoid potential microbial colonization of the rhizosphere. This eliminated any potential microbial deposition of metal oxyhydroxide-plaques (e.g. Fe and Mn) on the root epidermal surface (29, 30) known to reduce the uptake of other metals such as Cu, Mg, Ni, and P (31-34). Europium(III) chloride addition (0.001-0.01 M) to the growth media did not cause any immediate phytotoxic symptomology to either the roots or shoots. Slight tip burn and interveinal chlorosis were evident after 5-7 days of 0.01 M Eu(III) exposure. Therefore, all spectroscopic measurements were performed within the first 4 days following exposure. Microscopically, within 2-3 h following exposure a dark spot appeared in the root apical meristem just behind the root cap. The presence of Eu(III) in the dark spot was identified by its fluorescence characteristics to be described in the next section. Over time, the darkened region gradually extended along the entire root structure within the steele, although the intensity of the darkened region reduced dramatically beyond the root tip meristem. Because of the root’s translucent nature, we found that by placing the plant in the custom-built cuvette (Figure 1b), both optical and fluorescence microscopic measurements could be collected on a live root at the same time. Further with the microscope in a confocal configuration, both optical and fluorescence measurements could be collected with high spatial (1 µm) resolution. Fluorescence Characteristics of Eu(III) Assimilation by Roots. Both the oat root and Eu(III) are fluorescent. Figure 2 shows the steady-state fluorescence emission spectra of an VOL. 37, NO. 22, 2003 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 3. Steady-state fluorescence emission spectra of (a) Eu(III) accumulated by oat plant roots; (b) Eu(III) accumulated by oat plant root and transported to oat plant leaves; (c) 0.05 M EuCl3 aqueous solution; (d) 0.01 M EuCl3 in oat plant growth media. The broad background fluorescence of the plant root have been corrected. λex ) 394 nm. oat root which was rinsed with DDI water (trace a) and the growth media (trace b) prior to exposure to Eu(III) as well as 0.01 M aqueous EuCl3 solution (trace c), excited at 394 nm, corresponding to the absorption maximum of Eu(III) (14). The emission spectra of oat roots are intense and broad with emission maximum located at ∼460 nm. The appearance of a shoulder band at 440 nm suggested that more than one type of fluorophores were likely present in the root. A detailed identification of the spectral origin of these broad fluorescence emission is beyond the scope of this work. However, it has been generally accepted that such blue fluorescence is related to the phenylpropanoid type metabolites from plant roots, such as ferulic acid and p-coumaric acid (35, 36). The steady-state emission spectra of aqueous Eu(III) consisted of a set of sharp bands with emission maximum located at 592 nm (Figure 2, trace c). The origin of these sharp emission bands has been well-documented for aqueous Eu(III) and its complexes (16, 37). The ground state of Eu(III) is 7F0, and the lowest excited state is 5D0. The weak emission bands between 445 and 560 nm correspond to the radiative transitions from the higher excited states (5D2, 5D1) to the ground state while the five bands at 575, 592, 616, 640, and 700 nm correspond to the transitions from the lowest excited state (5D0) to the ground state (7Fj, j) 1, 2, ..., 4), respectively. The 5D0 f 7F1 transition (ca. 592 nm) has a strong magnetic dipole character, and thus its radiative transition probability is minimally affected by the ligand electric field. In contrast, the 5D0 f 7F2 has a predominantly electric dipole character, and its radiative transition probability is very sensitive to the ligand environment. It has been shown that complexation with chelating ligands such as β-diketones and polyaminopolycarboxylic acids cause both a shift of the band position toward the longer wavelength and a significant increase of the spectral intensity (14, 16, 38). Three days after introduction of 0.01 M EuCl3 into the growth media, the plant was removed from the growth media, and the roots were thoroughly rinsed with DDI water. Steadystate fluorescence emission spectra were recorded for both the roots and leaves of the same oat plant via excitation at 394 nm. The resulting emission spectra consisted of sharp Eu(III) emission bands overlapped on the broad emission from the plant itself. The profile of the broad fluorescence emission from the plant root alone did not change with, or without, the presence of Eu(III); hence fluorescence spectra of assimilated Eu(III) were obtained by subtracting the background fluorescence spectra of a control (minus Eu(III)) plant. The resulting emission spectra of plant accumulated Eu(III) are shown in Figure 3 along with the emission spectra of Eu(III) chloride in both DDI water and the growth media in the same wavelength range. 5250

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FIGURE 4. Fluorescence decay data and its best-fit curve of (a) oat plant alone, λex ) 300 nm, λem ) 460 nm; (b) Eu(III) assimilation in oat plant roots; (c) Eu(III) in oat plant growth media. In (b) and (c), λex ) 394 nm, λem ) 616 nm. The fluorescence emission spectra of assimilated Eu(III) in both the oat roots and leaves (Figure 3, traces a and b) show distinctive difference from those of Eu(III) in DDI water and the growth media (Figure 3, traces c and d). First, the relative intensity of the 616 nm band to that of the 592 nm band increases significantly. In DDI water and the growth media the fluorescence intensity at 616 nm was only 25% of that at 592 nm. However, for assimilated Eu(III) the fluorescence intensity at 616 nm is 50% more than that at 592 nm. Second, a small but noticeable red-shift of the 616 nm band is observed for the accumulated Eu(III) relative to aqueous Eu(III). These results are similar to the spectral changes observed when Eu(III) is complexed to carboxylic acids and aminocarboxylic acids (14, 38) in which the waters of hydration are replaced. Therefore, a strong inner-sphere complex is likely to have formed between the assimilated Eu(III) cation and the cellular components of the oat root. The similar Eu(III) spectra recorded for the leaves of the same plant suggests that the Eu(III) assimilated by the oat root was transported to the leaves and that the same type of complexes were formed. However, from these data it is unclear if the form of the translocated Eu(III) was as the ion itself or in a Eu-ligand form. On the other hand, the emission spectra of Eu(III) in the growth media resembled that of aqueous Eu(III) chloride. This suggests that, under the experimental conditions, Eu(III) does not, or may only form very weak complexes with plant constituents potentially secreted by oat root into the growth media, or that the relative free Eu(III) concentration to the complex form was so high as to obscure any changes. Plants can release carboxylic acids into the hydrosphere (39). If the plant had released such chemicals to complex and mobilize metal ions as Eu(III) in order to facilitate metal uptake, the binding characteristics between Eu(III) and these materials would still be entirely different from those between the assimilated Eu(III) and the plant. Fluorescence lifetime of the oat roots measured by excitation with picosecond laser pulses (λex ) 300 nm) indicated that the broad fluorescence (λmax ) 440 or 460 nm) decays biexponentially (Figure 4, trace a) with fluorescence lifetimes of 1 and 10 ns, respectively. Such short fluorescence lifetimes are typical of the organic and biological fluorophores (40). The appearance of two decay components is consistent with the presence of two major types of fluorophores evidenced by overlapping fluorescence emission spectra with emission maxima at 440 and 460 nm, respectively. However, after the oat roots were exposed to EuCl3, an additional fluorescence decay component with much longer lifetime (0.345 ms) was observed (Figure 4, trace B) when excited with nanosecond laser pulses (λex ) 394 nm). Fluorescence decay curves recorded at different locations along the root

showed little variation of the fluorescence decay constant. Weak fluorescence emission of Eu(III) in oat leaves did not allow for an accurate analysis of fluorescence decay curves. Under the same conditions, the fluorescence of Eu(III) in the growth media exhibited a single exponential with a fluorescence lifetime of 0.117 ms, the same as that of uncomplexed Eu(III) in aqueous solutions (14). The relatively longer Eu(III) lifetime, 0.345 ms, as compared with that in the growth media (0.117 ms) suggests that the accumulated Eu(III) exists as a species statistically different than the hydrated cation, Eu3+(aq). It has been shown that there is a linear relationship between the fluorescence decay constant and the number of water molecules remaining in the inner sphere of Eu(III) ion (19)

nH2O ) 1.07‚kH2O - 0.7

(1)

where kH2O is the measured fluorescence decay constant of Eu(III) (in ms-1) and nH2O is the number of water molecules remaining in the inner coordination sphere of Eu(III). Substituting the measured fluorescence decay constant into eq 1 yields an nH2O for assimilated Eu(III) of 2.8 ( 0.5. Assuming a nanohydration number for aquous Eu(III) (15), a hydration number of 2.8 suggests six water molecules in the inner sphere of Eu(III) are replaced upon complexation with the oat root material. In other words, there are up to six donor atoms from the plant root bind to each assimilated Eu(III) cation. The chemical composition and structure inside a plant root is complex at the molecular level. The cellular constituents of a plant root include a variety of functional groups such as amino acids, proteins, nucleic acids, and phosphate as well as other phenolic, hydroxyl, thioether, and amide moieties, etc. (20, 41, 42). Ke and Rayson studied the binding of uranyl to isolated root cells of Datura innoxia by fluorescence emission and lifetime measurement at liquid nitrogen temperature (41). By comparing the fluorescence decay characteristics of uranyl sorbed in Datura innoxia and uranyl in several model solutions containing carboxyl, amine, hydroxyl, phosphoryl, sulfate, and sulfonate functionalities, they identified phosphoryl and dicarboxyl groups as the dominant functional groups responsible for the binding of uranyl to Datura innoxia cell walls. Fluorescence spectroscopic studies of Eu(III), Cu(II), Gd(III), and Cd(II) binding to Datura innoxia cell walls indicates that there are two types of totally different chemical moeities containing two different kinds of carboxylate groups (20). One contains a single carboxylate group which binds to the metal ion in a 1:1 ratio, while the second contains two carboxylate groups which binds in a 2:1 ratio and has a greater binding strength than the former. In contrast, EXAFS and XANES studies of Au+ binding to Chlorella vulgaris primarily occurred at the sites containing sulfur and nitrogen as the ligating atoms (43). Similarly, Gardea-Torresdey et al. (44) found that gold(III) binding capacities increased slightly, while those of Cu(II) and Al(III) decreased upon esterification of carboxylate groups associated with the surface of five different algal species. Europium(III), being a hard acid like UO22+ and Al3+ (45), should exhibit preferential binding to carboxylate, phosphate, and aminocarboxylate groups inside the root structure. At present the exact nature of the Eu(III)-oat root binding, distribution and characteristics is unclear. The long lifetimes observed in the present study suggests that accumulated Eu(III) in oat roots is not due to deposition of its hydrolysis products since the short internuclear distance typical of hydrolysis products between adjacent Eu(III) atoms causes fluorescence quenching due to inter-Eu(III) energy transfer, resulting in considerably shorter fluorescence lifetimes (46, 47).

FIGURE 5. Relative fluorescence intensity of Eu(III) along the midline axis of oat plant root measured with a confocal fluorescence microscope. λex ) 394 nm, λem ) 616 nm following 120-h exposure to 10 mM Eu(III). Eu(III) Distribution Inside Oat Root and Uptake Rate. Histologically, Eu(III) fluorescense was observed to decrease away from the root tip along the root axis (Figure 5). The highest intensities were always found in the apical meristematic tissue below the root cap at the root tip and for up to 300-500 µm basipetally. This confirmed that earlier observations of the dark spots formed around the root cap following exposure of the roots to EuCl3 solution were due to the intake of Eu(III). The darkness of the spots were proportional to the intensity of Eu(III) fluorescence. It is hypothesized that Eu(III) binding to the nucleic acids (nuclear material) in these cells leads to its accumulation within this tissue. The apical meristem is a region of active cell division, and the ratio of nucleic acids and proteins to cell volume is very high. Fast diffusion may also play an important role since the effective porosity will be higher in a region of active cell division. This is consistent with the observation by Hardiman (39, 48) that most heavy metal uptake was performed by younger parts of the root since in this part of the root the Casparian strips are not fully developed and there is a higher likelihood that metal ions can diffuse into the xylem vessels, thus facilitating faster transport. Another common observation of elevated fluoresence was found distally from the meristem within the zone of maturation. The developing epidermal root hairs of this region exhibited fluorescence within their cytoplasm and nuclei. This would be expected given the function of these structures in the uptake of potassium and phosphorus (39) from the soil solution. Transverse scans were made across the intact roots just basipetal to the zone of maturation. A rapid rise in fluorescence was observed at the epidermis. This was followed by a slight decline throughout the cortex and another rise through the vascular cylinder of the stele (Figure 6). Clearly, there is differential distribution of assimilated Eu(III) within the root. To correlate the concentration of accumulated Eu(III) with different root tissues and cell types, free-hand cross sections approximately 300-500 µm thick were taken from the same roots and also subjected to transverse scans. Immediately following the scanning the sections were stained with 0.1% (w/v) toluidine-O blue dye, rinsed in distilled H2O, and photographed, allowing identification of root tissue/ cell type at the measured location (Figure 7). The relative fluorescence intensity (Figure 7) shows that the epidermis, exposed to the Eu in the medium, is elevated in concentration. This is likely due to diffusional uptake as opposed to physical deposition (49) as suggested from the long fluorecsence lifetimes. The apparent decrease of Eu(III) VOL. 37, NO. 22, 2003 / ENVIRONMENTAL SCIENCE & TECHNOLOGY

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FIGURE 6. Relative fluorescence intensity of Eu(III) along a cross section of an oat plant root, ∼1162 µm away from the root tip measured with a confocal fluorescence microscope. λex ) 394 nm, λem ) 616 nm.

sorption initially occurs at the meristem near the root cap and transports upward in the root while Eu(III) diffusion across the epidermis and cortical tissue into the stele is limited. The results of this study are significant in that they reflect real time metal uptake and assimilation/transport in a living tissue, in this case the growing root of an intact transpiring plant. The distributions of the metal among the various cell types of the root confirm some of the previous data with killed and fixed tissues but can provide insight into the actual chemical state of the metals within the living cells. Applications of this technique with even more fluorescent metal probes, such as Cm(III) and U(VI) which allow spectroscopic measurements at much lower metal concentrations, e.g. 10-9 M (54-56), may provide molecular-level information that will aid in the determination of efficient varieties for phytoremediation and actual pathways for pollutants (heavy metals/radionuclides) within the plant rhizosphere as well as approaches for questions on basic mineral nutrition of plants.

Acknowledgments This research was supported by the Rhizosphere Microbial Ecology Laboratory Directed Research and Development project at the Pacific Northwest National Laboratory (PNNL) and performed at the W.R. Wiley Environmental Molecular Sciences Laboratory, a national scientific user facility sponsored by the Department of Energy’s Office of Biological and Environmental Research and located at PNNL. Pacific Northwest National Laboratory is operated for the U.S. Department of Energy by Battelle under Contract DE-AC0676RLO 1830.

Literature Cited

FIGURE 7. Relative fluorescence intensity of Eu(III) in different tissues of an oat plant along a cross section measured with a confocal fluorescence microscope. λex ) 394 nm, λem ) 616 nm. EPID ) epidermis; END ) endodermis; XP ) xylem parenchyma; SCC ) sieve element-companion cell complex; PX ) protoxylem. in the cortical tissue suggests that diffusion of Eu(III) across the large cortical cells toward the stele was not the favored transport mechanism of Eu(III) (50). At the endodermis the fluorescence level again rise suggesting deposition or accumulation here at the region of the developing Casparian bands (51). Within the stele the highest concentration was found within the developing xylem parenchyma, cells reported to control composition of the transpiration stream (52), and the companion cells of the phloem, cells known to have high concentrations of ribosomes and therefore high nucleic acid levels (53). Lower concentrations were found within the lumens of the protoxylem and developing metaxylem which indicates that the assimilated Eu(III) may either have been leached out to the medium following sectioning or that the concentration has been diluted within the transpiration stream. For the physiologically dependent nutrients such as Zn, Cu, and iron, it was found that the transfer of these metal ions occurs mainly through the xylem and phloem (39). The relatively high Eu(III) in the stele of oat root suggest that for heavy metal ions, such as Eu3+, its transport follows the same routes. This is consistent with the observation that under relatively high Eu(III) concentration in the media Eu(III) 5252

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(1) Chen, H.; Cutright, T. Chemisphere 2001, 45, 21-28. (2) Garbisu, C.; Alkorta, I. Biosour. Technol. 2001, 77, 229-236. (3) Raskin, I.; Smith, R. D. Curr. Opin. Biotechnol. 1997, 8, 221226. (4) Vesk, P. A.; Allaway, W. G. Aquat. Bot. 1997, 59, 33-44. (5) Zararsiz, A.; Kirmaz, R. J. Radioanal. Nucl. Chem. 1997, 222, 257-262. (6) Cavallini, A.; Natali, L.; Durante, M.; Maserti, B. Sci. Tot. Environ. 1999, 243/244, 119-127. (7) Vesk, P. A.; Nockolds, C. E.; Allaway, W. G. Plant Cell Environ. 1999, 22. (8) Wood, R. M.; Patrick, J. W.; Offler, C. E. Ann. Bot. 1994, 73, 151-160. (9) Drew, M. C.; Webb, J.; Saker, L. R. J. Exp. Bot. 1990, 41, 815-825. (10) Gilroy, S. Annu. Rev. Plant Physiol. Plant Mol Biol 1997, 48, 165-190. (11) Knoblauch, M.; van Bel, A. J. E. Plant Cell 1998, 10, 35-50. (12) Oparka, K. J.; Duckett, C. M.; Prior, D. A. M.; Fisher, D. B. Plant J. 1994, 6, 759-766. (13) Fricker, M. D.; Oparka, K. J. J. Exp. Bot. 1999, 50, 1089-1100. (14) Wang, Z.; van de Burgt, L. J.; Choppin, G. R. Inorg. Chim. Acta 1999, 293, 167-177. (15) Choppin, G. R. Lanthanide Probes in Life, Chemical and Earth Sciences: Theory and Practice; Elsevier: New York, 1989. (16) Richardson, F. S. Chem. Rev. 1982, 82, 541-552. (17) Horrocks, W. D., Jr.; Albin, M. Progress in Inorganic Chemistry; John Wiley & Sons: New York, 1984; Vol. 31. (18) Horrocks, W. J.; Sudnick, D. R. J. Am. Chem. Soc. 1979, 101, 334-340. (19) Barthelemy, P. P.; Choppin, G. R. Inorg. Chem. 1989, 28, 33543357. (20) Ke, H.-Y. D.; Rayson, G. D. Environ. Sci. Technol. 1993, 27, 24662471. (21) Aruguete, D. M.; Aldstadt, J. H.; Mueller, G. M. Sci. Total Environ. 1998, 224, 43-56. (22) Jackson, L. J.; Rowan, D. J.; Cornett, R. J.; Kalff, J. Can. J. Fish Aquat. Sci. 1994, 51, 1769-1773. (23) Wyttenbach, A.; Tobler, L.; Furrer, V. Proc. 4th. Int. Conf. Biogeochem. Trace Elements, Berkeley, CA June 23-26, 1997 1997, 299.

(24) Liu, Z. The Effects of rare earth elements on growth of crops. In Proceedings of the Int. Symp. on New Results in the Research of Hardly Known Trace Elements and Their Role In Food Chain; Pais, 1988. (25) Holtom, G. R. SPIE 1990, 2-12. (26) Wang, Z.; Friedrich, D. M.; Ainsworth, C. C.; Beversluis, M. R.; Joly, A. G. J. Phys. Chem. A 2001, 942-950. (27) Foster, R. C.; Rovira, A. D.; Cock, T. W. Ultrastructure of the Root-Soil Interface; The American Phytopathological Society: St. Paul, MN, U.S.A., 1983. (28) Crowley, D. E.; Zdenko, R. Food Prod. Press 1999, 1-40. (29) Crowder, A. A.; Coltman, D. W. J. Plant Nutr. 1993, 16, 589-599. (30) Emerson, D.; Weiss, J.; Megonigal, J. Appl. Environ. Microbiol. 1999, 65, 2758-2761. (31) Greipsson, S. J. Plant Nutr. 1995, 18, 1659-1665. (32) Ye, Z. H.; Baker, A. J. M.; Wong, M. H.; Willis, A. J. New Phytologist 1997, 136, 469-480. (33) Christensen, K. K.; Sand-Jensen, K. Can. J. Bot.-Revue Can. Bot. 1998, 76, 2158-2163. (34) Batty, L. C.; Baker, A. J. M.; Wheeler, B. D.; Curtis, C. D. Ann. Bot. 2000, 86, 647-653. (35) Yamamoto, E.; Bokelman, G. H.; Lewis, N. G. 1988, 68-88. (36) Hartley, R. D.; Ford, C. W. J. Sci. Food Agric. 1989, 46(3), 301310. (37) Carnall, W. T.; Fields, P. R.; Rajnak, K. J. Chem. Phys. 1968, 49, 4450-4455. (38) Wang, Z.; Choppin, G. R.; Di Bernardo, P.; Zanonato, P. L.; Portanova, R.; Tolazzi, M. J. Chem. Soc., Dalton Trans. 1993, 2791. (39) Marschner, H. Mineral Nutrition of Higher Plants, 2nd ed.; Academic Press: New York, 1995. (40) Lackowicz, J. R. Topics in Fluorescence Spectroscopy; Plenum Press: New York, 1991. (41) Ke, H.-Y. D.; Anderson, W. L.; Moncrief, R. M.; Rayson, G. D. Environ. Sci. Technol. 1994, 28, 586-591.

(42) Ke, H.-Y. D.; Birnbaum, E. R.; Darnall, D. W.; Rayson, G. D. Environ. Sci. Technol. 1992, 26, 782-788. (43) Watkins, I. J. W.; Elder, R. C.; Greene, B.; Darnall, D. W. Inorg. Chem. 1987, 26, 1147-1151. (44) Gardea-Torresdey, J. L.; Becker-Hapak, M. K.; Hosea, J. M.; Darnall, D. W. Environ. Sci. Technol. 1990, 24, 1372-1378. (45) Pearson, R. G. J. Am. Chem. Soc. 1963, 85, 3533. (46) Blasse, G. Prog. Solid State Chem. 1988, 18, 79-171. (47) Takahashi, Y.; Kimura, T.; Minai, Y. Geochim. Cosmochim. Acta 2002, 66, 1-12. (48) Blasse, G.; Brxner, L. H. J. Solid State Inorg. Chem. 1989, 26, 367-382. (49) Kelly, C.; Mielke, R. E.; Dimaquibo, D.; Curtis, A. J.; Dewitt, J. G. Environ. Sci. Technol. 1999, 33, 1439-1443. (50) Steudle, E.; Peterson, C. A. J. Exp. Bot. 1998, 49, 775-788. (51) Steudle, E. J. Exp. Bot. 2000, 51, 1531-1542. (52) Boer, A. H. d.; Wegner, L. H. J. Exp. Bot. 1997, 48, 441-449. (53) Oparka, K. J. Annu. Rev. Plant Physiol. Plant Mol. Biol. 2000, 51, 323-347. (54) Wang, Z.; Zachara, J.; Felmy, A. R.; Liu, C.; Qafoku, O.; Catalano, J. Fluorescence Spectroscopic Studies of Uranium-Bearing Vadoze Zone Sediments. In Digest of Science and Technology Program Evaluation; Pacific Northwest National Laboratory: 2003. (55) Wang, Z.; Felmy, A. R.; Xia, Y. X.; Mason, M. J. Radiochim. Acta 2003, 91, 329-337. (56) Moulin, C.; Decambox, P.; Mauchien, P. J. Radioanal. Nucl. Chem. 1997, 226, 135-138.

Received for review April 16, 2003. Revised manuscript received September 4, 2003. Accepted September 10, 2003. ES0343609

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