Sedum alfredii is one of a few species known to hyperaccumulate zinc (Zn) and cadmium (Cd). Xylem transport and phloem remobilization of Zn in hyperaccumulating (HP) and nonhyperaccumulating (NHP) populations of S. alfredii were compared.
Micro-X-ray fluorescence (μ-XRF) images of Zn in the roots of the two S. alfredii populations suggested an efficient xylem loading of Zn in HP S. alfredii, confirmed by the seven-fold higher Zn concentrations detected in the xylem sap collected from HP, when compared with NHP, populations. Zn was predominantly transported as aqueous Zn (> 55.9%), with the remaining proportion (36.7–42.3%) associated with the predominant organic acid, citric acid, in the xylem sap of HP S. alfredii.
The stable isotope 68Zn was used to trace Zn remobilization from mature leaves to new growing leaves for both populations. Remobilization of 68Zn was seven-fold higher in HP than in NHP S. alfredii. Subsequent analysis by μ-XRF, combined with LA-ICPMS (laser ablation-inductively coupled plasma mass spectrometry), confirmed the enhanced ability of HP S. alfredii to remobilize Zn and to preferentially distribute the metal to mesophyll cells surrounding phloem in the new leaves.
The results suggest that Zn hyperaccumulation by HP S. alfredii is largely associated with enhanced xylem transport and phloem remobilization of the metal. To our knowledge, this report is the first to reveal enhanced remobilization of metal by phloem transport in hyperaccumulators.
If you can't find a tool you're looking for, please click the link at the top of the page to "Go to old article view". Alternatively, view our Knowledge Base articles for additional help. Your feedback is important to us, so please let us know if you have comments or ideas for improvement.
Elevated concentrations of heavy metals inhibit the metabolism of most plant species (Prasad & Hagemeyer, 1999). The metal hyperaccumulators have the rare ability to colonize heavy metal-polluted soils and to tolerate high heavy metal concentrations within plant tissues (Baker & Brooks, 1989; Baker et al., 2000). The elucidation of the mechanisms of metal hyperaccumulation may enable the rational design of technologies for the clean-up of metal-contaminated soils and for the bio-fortification of trace elements in crop food (McGrath & Zhao, 2003; Zhao & McGrath, 2009; Kramer, 2010).
Metal accumulation mechanisms in plants are generally controlled at three levels: uptake from the soil by the root system, translocation of metal ions to the shoot and the capacity for metal storage in leaves. In past decades, the majority of studies into the physiological and molecular mechanisms of metal uptake, transport, accumulation and tolerance in hyperaccumulator species have focused on members of the Brassica family, such as Noccaea caerulescens (formerly Thlaspi caerulescens) (Salt et al., 1999; Kupper et al., 2004; Callahan et al., 2007; Ebbs et al., 2009) and Arabidopsis halleri (Kupper et al., 2000; Zhao et al., 2000; Sarret et al., 2002, 2009; Hanikenne et al., 2008; Ueno et al., 2008; Gustin et al., 2009). A basic understanding of metal hyperaccumulation mechanisms has been elucidated, including enhanced metal uptake, increased xylem loading and increased detoxification in the shoot (Verbruggen et al., 2009). Several transporters involved in these physiological processes have been identified, such as AhHMA4 and TcHMA3 (Hanikenne et al., 2008; Ueno et al., 2011). Although considerable progress has been made, there is so far very little knowledge regarding the phloem transport processes in the metal hyperaccumulators.
Sedum alfredii Hance (Crassulaceae), found in contaminated soils of Zheijiang Province, China, has been identified as a zinc/cadmium (Zn/Cd) co-hyperaccumulator (Yang et al., 2002, 2004; Tian et al., 2009), and is one of only a few nonbrassica species to have demonstrated both Zn and Cd hyperaccumulation (Lu et al., 2008; Kramer, 2010; Claire-Lise & Nathalie, 2012). This plant species accumulates up to 2.9% Zn (dry weight, DW) in shoot without exhibiting toxicity symptoms (Yang et al., 2002; Tian et al., 2009). The mechanism of metal accumulation in this plant species is not fully understood. The aim of the present study was to investigate the characteristics of xylem transport and phloem remobilization of Zn in the hyperaccumulator S. alfredii (hyperaccumulating population, HP; formerly hyperaccumulating ecotype, HE) in comparison with its contrasting nonhyperaccumulating population (NHP, formerly nonhyperaccumulating ecotype, NHE). The in vivo distribution patterns of Zn and other elements in roots of HP and NHP S. alfredii were investigated by micro-X-ray fluorescence (μ-XRF) mapping. The concentrations of Zn and organic acids in the xylem sap collected from the two plant ecotypes were analyzed, with the Zn ligand environments in the xylem sap of the HP population determined using extended X-ray absorption fine structure (EXAFS). In addition, we used the stable isotope 68Zn as a tracer to study the characteristics of remobilization of Zn from old mature leaves to new growing young leaves in the two contrasting populations of S. alfredii. Subsequent analysis by μ-XRF mapping and laser ablation-inductively coupled plasma mass spectrometry (LA-ICPMS) provided additional insights into the cellular distribution patterns of total Zn and 68Zn in young leaves after remobilization, respectively.
Materials and Methods
Seedlings of two contrasting populations of Sedum alfredii Hance were cultivated hydroponically. Seeds of HP S. alfredii were obtained from an old lead (Pb)/Zn mine area in Qvzhou, and those of NHP S. alfredii were obtained from a tea plantation in Hangzhou, both in Zhejiang Province, China. The seeds of the two populations were germinated on a mixture of perlite and vermiculite moistened with deionized water. Four weeks after germination, the plants were subject to 4 d of exposure to one-quarter and one-half strength nutrient solution containing 2.0 mM Ca2+, 4.0 mM , 1.6 mM K+, 0.1 mM , 0.5 mM Mg2+, 1.2 mM , 0.1 mM Cl−, 10 μM H3BO3, 0.5 μM MnSO4, 5.0 μM ZnSO4, 0.2 μM CuSO4, 0.01 μM (NH4)6Mo7O24 and 100 μM Fe-EDTA. The nutrient solution pH was adjusted daily to 5.8 with 0.1 N NaOH or HCl. Plants were afterwards cultured in full-strength nutrient solution, which was continuously aerated and renewed every 3 d. Plants were grown in a growth chamber in a 16 h : 8 h photoperiod at 400 μmol m−2 s−1, day : night temperature of 26 : 20°C and day : night humidity of 70 : 85%.
Zn and organic acid in xylem sap
Seedlings of HP and NHP S. alfredii precultured in full nutrient solution for 8 wk were used for xylem sap collection. The nutrient solutions were replaced with fresh solutions containing different Zn concentrations 8 h before the onset of xylem sap collection. Treatments included 5 (control), 25, 100, 250 and 500 μM, each with four replicates. Twelve plants in one culture vessel were treated as one replicate. Xylem sap was collected according to Lu et al. (2008). Briefly, plants were de-topped using sharp blades at c. 3.0 cm above the junction point of root and shoot. Immediately after de-topping, each stem was rinsed with deionized water and blotted with absorbent paper. After discarding c. 0.3 ml of sap, each cut surface was blotted again and silicon tubing was fitted over the stem. Sap flowing from the tubing was collected in sterile vials for 1 h. At the end of the collection period, sap samples collected from the 12 plants were pooled to give one pool per container. Half of the xylem sap was immediately frozen at −20°C for elemental and organic acid determination, and the other was added to 30% glycerol and frozen at −80°C for EXAFS analysis. Roots were desorbed in 1.0 mM EDTA for 5 min, rinsed in deionized water, oven dried at 65°C for 72 h, weighed and ground using a stainless steel mill to pass a 0.25-mm sieve. Dry root samples (0.1 g) were digested with 5.0 ml HNO3–HClO4 (4 : 1, v/v), and the digest was transferred to a 50-ml volumetric flask, made up to volume and filtered. For xylem sap samples, a subsample of 0.5 ml was mixed with 25 ml of 2% (w/v) nitric acid (Lu et al., 2008). Zn concentrations of the samples were determined by inductively coupled plasma-atomic emission spectroscopy (ICP-AES) (Shimadzu ICPS-7510; Shimadzu, Tokyo, Japan). Organic acids in the collected xylem sap were analyzed using high-performance liquid chromatography (HPLC) (Shimadzu LC-10ATvp) with an ion-exchange analytical column C18 (5 μm, 250 mm × 4.6 mm) and an eluent of 15 mM KH2PO4 (pH 2.5), according to Yang et al. (2006). Organic acids were detected by a UV detector (Shimadzu SPD-10ATvp) at 205 nm.
EXAFS analysis of Zn speciation in xylem sap
EXAFS data of xylem sap from the HP S. alfredii plants were collected at the Stanford Synchrotron Radiation Laboratory (SSRL) with the storage ring SPEAR-3 operating at 3.0 GeV and with ring currents of 80–100 mA. Zn K-edge spectra were recorded on beamline 7-3, SSRL, with an upstream rhodium (Rh)-coated collimating mirror, a silicon (Si) (220) double-crystal monochromator and a downstream focusing mirror. The incident X-ray intensity was monitored using a krypton (Kr)-filled ionization chamber. The monochromator energy of each spectrum was calibrated using Zn metal foil between the second and third ionization chambers; its absorption edge was calibrated to an edge of 9659 eV. Zn Kα fluorescence was recorded using a 30-element germanium detector (Canberra Industries, Meriden, CT, USA) equipped with Soller slits and copper (Cu) filters. During data collection, samples were maintained at approximately 10 K in a liquid helium flow cryostat to minimize the loss of intensity in the signal. Spectra were also collected for standard Zn species, including aqueous Zn (Zn(NO3)2), Zn-citrate (solution), Zn-malate (solution), Zn-oxalate (solution), Zn-phytate (solution), Zn-histidine (solution), Zn-succinate (solution), Zn-glutathione (solution) and Zn-cysteine (solution). All reference solutions and xylem sap were prepared in 30% glycerol to prevent ice crystal formation. The complexes of Zn-citrate, Zn-malate, Zn-oxalate, Zn-phytate, Zn-histidine, Zn-succinate, Zn-glutathione and Zn-cysteine were made by adding 5.0 mM citrate, malate, oxalate, phytate, histidine, succinate, glutathione and cysteine to an aqueous solution of 0.5 mM Zn(NO3)2, pH 6.0. Spectra of all plant samples and standard solution samples were recorded in fluorescence mode, whereas spectra of solid standard samples were recorded in transmission mode. Detuning of the primary beam by 50% was performed to reject higher harmonic reflections.
Sixteen scans were collected and averaged for each sample to improve the signal-to-noise ratio. The spectra data of each sample were calibrated to the Zn K-edge (9659 eV) in the program SIXpack (Webb, 2005). The normalization of the EXAFS spectra was carried out according to standard methods using the SIXpack program suite. The spectra were normalized to unit step height using a linear pre- and post-edge background subtraction, and transformed to k-space based on E0 equal to 9659 eV. The k-function was extracted from the raw data by subtracting the atomic background using a cubic-spline consisting of seven knots set at equal distances fitted to k3-weighted data; k3-weighted χ(k)-functions were Fourier transformed over the 1.5–12-Å−1 range using a Kaiser–Bessel window with a smoothing parameter of 4. The k3-weighted EXAFS spectra recorded on the plants were least-squares fitted over a wave vector (k) range of 1.5–12 Å−1 using a combination of Zn compounds as standard. Best fits were derived by incrementally increasing the number of fitted components and minimizing the fitted residual. The range for the fit was varied as a function of data quality and in order to test contributions from minor components. In parallel, the structural parameters for the second Zn coordination shells were determined with simulation using SIXpack (Webb, 2005). The EXAFS spectra were Fourier transformed over a k range of 3–12 Å−1, and the contribution of the first and/or second shell was simulated in k- and R-space. Phase and amplitude fractions were calculated by FEFF8 ab initio code for Zn–O and Zn–C pairs, Zn histidine dehydrate for Zn–N and Zn–C pairs, and hopeite for Zn–O, Zn–P and Zn–Zn pairs.
Redistribution of 68Zn applied to mature leaves
The 68Zn solution was prepared according to Wu et al. (2010) with some modifications. 68Zn-enriched isotope was purchased from Isotec Inc. (Miamisbug, OH, USA). The isotopic composition of enriched isotope was as follows (atom%): 64Zn, 2.45; 66Zn, 1.45; 67Zn, 0.73; 68Zn, 95.1; 70Zn, 0.22. The natural isotopic composition of Zn nutrient solution is as follows (atom%): 64Zn, 48.63; 66Zn, 27.90; 67Zn, 4.10; 68Zn, 18.75; 70Zn, 0.62. A known amount of the enriched preparation of 68Zn was dissolved in a few drops of 1 N acetic acid with shaking, followed by an equivalent volume of 0.1 N H2SO4. The sample was heated at 75°C to near dryness and then rediluted to the desired volume with double-deionized (DDI) water. The pH of the solution was adjusted to 5.8 with 0.5 N KOH.
Plants of the two populations of S. alfredii were precultured in full nutrition solution for 4 wk. Before 68Zn treatment, all the plants were exposed to nutrient solution without Zn for 3 d, and then six mature leaves from each plant were washed thoroughly with deionized water and treated with 5.0 ml of 2.0 mM 68ZnSO4 solution (pH 5.8) containing 0.01 Silwet L-77 spray adjuvant (v/v); mature leaves of control plants received the same treatments, but without 68ZnSO4 in the labeling solution. Mature leaves were soaked in the solution for 10 s; other plant tissues were protected by a Teflon membrane from contamination. Four plants were treated as one replication, and each treatment had three replications. The foliar application of 68Zn was replicated one time per day for 7 d, and the plants were harvested after 4 d. During the treatments, the plants were constantly exposed to nutrient solution without Zn. The concentrations of 68Zn and total Zn in the new growing leaves of collected 68Zn-treated plants and controls were determined by ICPMS (Agilent 7500a; Agilent Technologies, Palo Alto, CA, USA) and ICP-AES (Shimadzu ICPS-7510). The amount of 68Zn in the new growing leaves derived from foliar treatment was calculated from the equation based on the isotope ratio modified from Wu et al. (2010).
New growing leaves were cut from the plants treated with 68Zn and were then surface rinsed. Leaf samples at similar developmental stages from treated and untreated plants were selected from both populations of S. alfredii for comparisons. Leaf cross-sections (100 μm thick) were cut with a cryotome (CM1850; Leica, Nussloch, Germany) at a temperature of −20°C (Tian et al., 2010). Sections in good condition were selected and freeze–dried at −20°C for 3 d. In a separate experiment, plants of the two populations of S. alfredii were precultured in full nutrition solution for 4 wk; fresh roots were collected from the plants after 100 μM Zn exposure for 24 h, rinsed, immediately frozen with liquid nitrogen and then freeze–dried at −20°C for 3 d.
The μ-XRF analysis of Zn and other elements of the plant samples was carried out on SSRL beamline 2-3 using an Si (220) double-crystal monochromator with harmonic rejection achieved by detuning one monochromator crystal to 50% peak intensity. The equipment conditions were the same as those in our previous report (Tian et al., 2010). The brilliance of the beam in beamline 2-3 was 1 × 1012 photons s−1 mm−2 mrad−2/0.1%of the bandwidth (BW). The microfocused beam of 2 μm was provided by a Kirkpatrick–Baez mirror pair (Xradia Inc., Pleasanton, CA, USA) with the sample at 45° to the incident X-ray beam. The fluorescence yield was detected using a single-channel Vortex Si detector (Hitachi High-Technologies Science America Inc., CA, USA). The elemental distribution maps were collected at room temperature using an X-ray beam of 5 μm pixel size. The dwell time per point was 200 ms. The beam energy was set to 15 keV during mapping. The fluorescence energies windowed for this investigation were phosphorus (P), sulfur (S), potassium (K), calcium (Ca), manganese (Mn), iron (Fe) and Zn. The maps were produced using the software package SMAK version 0.34, S-4 (http://www-ssrl.slac.stanford.edu/~swebb/smak.htm). The fluorescence data were presented as tricolor maps that allow for the spatial distribution of three elements to be shown. Pixel brightness was displayed in RGB (color model), with the brightest spots corresponding to the highest element fluorescence.
After μ-XRF analysis, the new growing leaf samples were subjected to 68Zn determination by the Interdisciplinary Center for Plasma Mass Spectrometry at the University of California at Davis (ICPMS.UCDavis.edu) using an Agilent 7500A ICP-MS (Agilent Technologies). Standard preparation and quantification, and elemental analysis of the samples determined by LA-ICPMS, were performed under the same optimized experimental parameters as described by Tian et al. (2011). Briefly, helium (0.85 l min−1) was used as the transport gas from the New Wave laser ablation cell (New Wave Research Inc., ESI, Fremont, CA, USA) and was mixed with argon gas (1.05 l min−1). The LA-ICPMS system was tuned for sensitivity before each experiment using the National Institute of Standards and Technology (NIST) 612 glass standard. The formation of oxides was minimized by monitoring the m/z 248/232 (232Th16O+/232Th+) ratio value at < 0.2%. Standard reference material (SRM) NIST 1570a spinach leaves was used for the standard calibration of the analytical data. The spinach leaves were dried at 75°C until constant weight was reached, and then ground into a fine powder. Half a gram of each standard material was weighed and pressed into a pellet without any binder under 8 atm pressure. The pelletized materials and plant samples were analyzed for elemental concentrations by LA-ICPMS. The ion intensities of 68Zn, 64Zn, 66Zn and 70Zn were recorded. The analytical procedures were validated by SRM NIST 1547 peach leaves and SRM NIST 1515 apple leaves. During the experiment, the spot size was 10 × 10 μm2. The scanning points of the samples were selected and observed using a microscope. Nine or ten replicates were performed for each of the scanning points in one specimen.
Statistical analysis of the data
The data on the concentrations of organic acids in the xylem sap (Table 1), total Zn and 68Zn in the new growing young leaves (Fig. 4) were analyzed statistically using the SPSS package (version 11.0; SPSS Inc., Chicago, IL, USA). Analysis of variance (ANOVA) was performed on the datasets, and the mean and SE of each treatment, as well as the least significant difference (LSD) (P <0.05 and P <0.01), for each set of corresponding data were calculated.
Table 1. Organic acid components in xylem sap of the two populations of Sedum alfredii under zinc (Zn) exposure
Zn levels (μM)
Concentration of organic acid (mg l−1)
Values represent means of four different replications (n =4). Different letters in the same column indicate significance of the treatments at P <0.05; one or two asterisks indicate significant difference between the two populations at P <0.05 or P <0.01, respectively. Populations: HP, hyperaccumulating; NHP, nonhyperaccumulating.
6.1 ± 2.7a
103.4 ± 15.1c**
44.8 ± 5.7a**
23.95 ± 3.71a
6.4 ± 1.6a
178.6 ± 14.8b**
31.5 ± 4.8b
22.56 ± 4.33a
7.0 ± 0.5a
235.8 ± 24.8a**
21.4 ± 4.6b
26.78 ± 2.74a
3.4 ± 1.9b
17.3 ± 7.0a
20.4 ± 4.2
6.3 ± 0.2a
15.2 ± 5.5a
8.4 ± 0.5a*
19.4 ± 1.2a
µ-XRF imaging of Zn in the roots of HP and NHP populations
The distribution patterns of Zn and other elements in the roots of HP and NHP S. alfredii were determined by μ-XRF mapping after 100 μM Zn exposure for 24 h (Fig. 1). The integrated intensities of Zn, K, Ca, Mn, Fe, Cu, P and S were calculated from X-ray fluorescence spectra and normalized by the intensity of the Compton scattering peak. Elemental mapping for the measurement area was obtained from the normalized intensity for each element. The distribution patterns of Zn (red), Ca (blue) and Fe (green) are presented in Fig. 1. Pixel brightness is displayed in RGB, with the brightest spots corresponding to the highest element fluorescence. Elemental maps for the other elements are not shown. Zn was preferentially distributed within the stele of NHP roots (Fig. 1b), whereas the distribution pattern of Zn in HP roots showed strong accumulation in the root epidermal cells (Fig. 1a). On the basis of the intensity of the Zn signal, the content of Zn in the root epidermis of HP plants was five-fold higher than that in the root stele (Fig. 1c), whereas Zn in the root stele was three-fold higher than that in the remainder of the roots in NHP plants (Fig. 1d).
Zn concentration in the xylem sap
Zn concentrations in the xylem sap of HP plants followed a nonsaturating curve with increasing Zn levels of 5–500 μM in the solution, whereas Zn concentration in the xylem sap of NHP plants increased linearly up to 100 μM and saturated above 100 μM, at least until 500 μM (Fig. 2a). Regardless of treatment, the Zn concentration in the xylem sap of HP plants was consistently much higher than that of NHP plants (P <0.01). At an external Zn supply of 25–500 μM, the Zn concentration in the xylem sap of HP plants was approximately seven- to nine-fold higher than that in the xylem sap of NHP plants (Fig. 2a). In contrast with the Zn concentration in xylem sap, there was no significant difference in Zn concentration in the roots of the two populations, although a slightly higher Zn concentration was observed in NHP plants at high Zn exposure levels (250–500 μM) (Fig. 2b).
Organic acids in xylem sap
In the xylem sap of HP S. alfredii, the predominant organic acids were citric acid, followed by malic acid, succinic acid and oxalic acid, whereas no succinic acid was observed in the NHP plant samples (Table 1). Fumaric acid, pyruvate, acetic acid and amber acid were not detected in the xylem saps of either population. In the control plants (5.0 μM Zn), the concentrations of citric acid in the xylem sap of HP S. alfredii were higher than 100 mg l−1, approximately six-fold greater than that in the xylem sap of NHP plants (Table 1). Exposure to high Zn levels (100 or 500 μM) increased significantly the concentration of citric acid in the xylem sap of HP plants, whereas no such effect was observed in NHP plants. High Zn treatments decreased the content of malic acid in the xylem sap of HP plants, but did not affect significantly succinic acid or oxalic acid.
Zn speciation in xylem sap
Bulk EXAFS was employed to investigate the Zn speciation in the xylem sap of HP S. alfredii treated with 100 μM or 500 μM Zn, which contained adequate metal concentrations to obtain good signal-to-noise ratios. The low concentrations of Zn in the xylem sap of NHP S. alfredii prevented the analysis of Zn speciation by this technique. Figure 3 shows Zn EXAFS spectra of xylem sap compared with model compounds, including aqueous Zn (Zn(NO3)2), Zn-citrate, Zn-malate, Zn-oxalate, Zn-phytate, Zn-histidine, Zn-succinate, Zn-glutathione and Zn-cysteine. The results of the EXAFS data, following refinement with SIXpack for the Zn ligand environment, in plant samples of HP S. alfredii are summarized in Table 2. Zn in the xylem sap from the plants of HP S. alfredii, treated with both 100 and 500 μM Zn, was exclusively coordinated with O ligands. The spectra were also analyzed using linear combination fitting (LCF) (Table 3), showing that a dominant chemical form of Zn in the xylem sap of HP S. alfredii was similar to aqueous Zn, with the remaining proportion of the metal associated as Zn-citrate.
Table 2. First and/or second shell structural parameters derived from the spectra of model compounds and xylem saps of hyperaccumulating (HP) Sedum alfredii plants treated with 100 and 500 μM Zn
σ2 (10−3 Å)
N, number of atoms; R, interatomic distance; σ2, Debye–Waller factor; R factor, reliability factor for goodness of fit, which was calculated as all spectra were recorded as frozen hydrate. SE, mathematical standard errors of the refinement (two sigma level).
6.2 ± 0.3
2.03 ± 0.03
6.3 ± 0.2
2.02 ± 0.04
3.9 ± 0.3
2.85 ± 0.02
5.9 ± 0.1
2.05 ± 0.03
2.5 ± 0.3
2.84 ± 0.02
5.4 ± 0.3
1.99 ± 0.04
2.2 ± 0.2
2.87 ± 0.03
4.1 ± 0.2
1.98 ± 0.01
1.0 ± 0.1
3.12 ± 0.02
4.0 ± 0.1
2.06 ± 0.01
3.5 ± 0.1
2.96 ± 0.05
6.3 ± 0.7
2.03 ± 0.04
4.0 ± 0.1
2.32 ± 0.01
3.9 ± 0.2
2.34 ± 0.02
100 μM Zn
5.4 ± 0.1
1.98 ± 0.02
4.4 ± 0.5
500 μM Zn
6.4 ± 0.3
2.12 ± 0.04
1.2 ± 1.0
Table 3. Zinc (Zn) species identified by linear combination fit (LCF) analyses of Sedum alfredii xylem sap spectra using a combination of Zn model compounds as standard
Zn levels (μM)
Zn species (%)
SE, mathematical standard errors of the refinement (two sigma level).
55.87 ± 7.43
6.07 ± 2.78
42.31 ± 4.54
64.12 ± 5.65
36.75 ± 6.32
Redistribution of Zn to the young leaf
To investigate the redistribution of Zn via the phloem, the mature leaves of HP and NHP S. alfredii were labeled with the stable isotope 68Zn for 7 d, and the concentrations of total Zn and 68Zn in the newly developed leaves were analyzed by ICP-AES and ICPMS, respectively. After 7 d of treatment, total Zn concentration in the HP plants reached 988 mg kg−1 DW in the newly developed leaves (Fig. 4a), which was four-fold higher than that in the unlabeled control plants. Application of 68Zn to NHP plants resulted in a significant, but much less pronounced, enrichment of Zn in newly developed young leaves (Fig. 4a). Across all treatments, total Zn concentrations in the newly developed leaves from the HP plants were significantly higher than those from the NHP plants (Fig. 4a). 68Zn in the newly developed leaves of HP plants treated with 68Zn averaged 789 mg kg−1 DW (Fig. 4b), demonstrating a rapid retranslocation of Zn from labeled mature leaves to young leaves. By calculating the difference between the 68Zn levels found in plant samples from 68Zn-labeled and unlabeled control plants, we were able to estimate the mass of 68Zn in the new growing leaves specifically retranslocated from the mature leaves. No significant difference in leaf growth was observed between treatments or populations over the duration of the labeling experiment. At the completion of the 7-d labeling experiment, 745 and 99 mg kg−1 DW of 68Zn was assimilated by the HP and NHP plants, respectively.
To investigate in more detail the cell layers to which Zn was redistributed from mature leaves to young leaves in the two populations of S. alfredii, the spatial distribution of Zn and other elements in the newly developed leaves was analyzed by μ-XRF mapping (Fig. 5). The Zn ion intensity in leaf cross-sections of the newly developed leaves from 68Zn-labeled HP plants showed a preferential localization to mesophyll cells surrounding the vascular bundles and in the epidermis (Fig. 5a). The intensity of the Zn signal was up to 150 counts s−1 in the mesophyll cells of HP plants (Fig. 5a), whereas only a very low Zn intensity was observed in the cross-sections of the newly developed leaves from NHP plants treated with 68Zn labeling (Fig. 5b). In the newly developed leaves from both HP and NHP control plants (without 68Zn labeling on the mature leaves), only a very low and diffuse Zn signal was observed (Supporting Information Fig. S1). No marked concentration of other elements (Ca, K, P, S, etc.) in the phloem environment was observed, and no significant correlation between the intensity of Zn and other elements was found (data not shown).
The newly developed leaves of HP plants treated with or without Zn labeling were further subjected to LA-ICPMS analysis of Zn isotopes (Fig. 6). There was no measurable localization of Zn (68Zn, 64Zn, 66Zn and 70Zn) in any cell type within the cross-sections of the newly developed leaves from the control plants of HP S. alfredii, with concentrations in the range 30–132 mg kg−1 DW. In the cross-sections of newly developed leaves from the HP plants treated with 68Zn labeling to the mature leaves, a clear preferential distribution of 68Zn was observed in the mesophyll tissues on either side of the vascular system, with concentrations up to 1167 mg kg−1 DW (scanning point 62). The enhanced concentration and localization were restricted to the 68Zn concentrations, with all other isotopes showing little localization of concentration.
An enhanced rate of root-to-shoot translocation is the pivotal process specifically expressed in hyperaccumulators (Lasat et al., 1996; Shen et al., 1997; Zhao et al., 2006; Lu et al., 2008). Efficient root-to-shoot translocation largely depends on several processes (Lasat et al., 1996, 1998): symplastic uptake by roots; root sequestration; xylem loading; and xylem unloading and uptake of metals by foliar cells. Our previous studies have revealed that reduced Zn sequestration in root cells (tonoplast) and stimulated Zn uptake in leaf cells are the important mechanisms involved in the strong Zn hyperaccumulation observed in S. alfredii (Yang et al., 2006). The results of this study suggest that the hyperaccumulation of Zn in S. alfredii is largely associated with efficient xylem loading. µ-XRF images of Zn in roots of S. alfredii showed that Zn was preferentially distributed within the root stele of NHP plants, whereas this distribution pattern of Zn was not observed for HP roots (Fig. 1). This contrast in Zn localization in roots for HP and NHP plants is probably the result of the efficient xylem loading of Zn in HP roots for subsequent translocation to the shoot, whereas Zn in the roots of NHP plants is less available for xylem loading, and thereby is concentrated around the root vascular cylinders. This hypothesis is confirmed by the seven-fold higher Zn concentrations detected in the xylem sap collected from HP relative to NHP plants (Fig. 2). Higher metal concentrations in the xylem sap as a result of enhanced xylem loading have been reported previously in hyperaccumulators (Lasat et al., 1998; Lu et al., 2008, 2009). Several types of transporter, such as P-type ATPase-HMA, are involved in the process of efficient xylem loading of metals in hyperaccumulators. In the Zn/Cd hyperaccumulator A. halleri, triplicate expression of AhHMA4 mediates the enhanced Zn loading into the xylem vessels necessary for shoot Zn hyperaccumulation (Hanikenne et al., 2008). The present study suggests that xylem loading in roots has an important role in the Zn hyperaccumulation process in S. alfredii. The genetic analysis of the transporters responsible for efficient xylem loading may help to facilitate a better understanding of Zn hyperaccumulation in this particular species.
The role of ligands in heavy metal transport in the xylem of hyperaccumulators is controversial. Several studies have reported that metals, such as Zn and Cd, occur mainly in the free ionic form during xylem transport by hyperaccumulators (Salt et al., 1999; Ueno et al., 2008), whereas others have suggested that metals are associated with certain ligands, such as histidine, citric acid or nicotianamine (Kramer et al., 1996; Kerkeb & Kramer, 2003; Durrett et al., 2007; Trampczynska et al., 2010). In the present study, the ligand environment of Zn in the hyperaccumulator S. alfredii was analyzed by EXAFS measurements. The results indicated that Zn was mostly transported as aqueous Zn, with the remaining proportion associated with citric acid in the xylem sap of HP S. alfredii. The association of Zn-citrate was supported by the extremely high concentrations of citric acid in the xylem sap of HP relative to NHP plants (Table 1). The citric acid concentrations in HP plants increased following Zn exposure, further implying an important role of citric acid in the xylem transport of Zn by HP S. alfredii. Evidence for the involvement of citrate in xylem transport for Zn exists from other species (Verbruggen et al., 2009). For example, a member of the MATE subfamily, FRD3, is thought to efflux citrate into the root vascular tissue, and this transporter is constitutively overexpressed in the hyperaccumulators A. halleri and N. caerulescens, and may play a role in Zn translocation (van de Mortel et al., 2006; Talke et al., 2006). LCF analysis of EXAFS spectra, however, is not free of uncertainties and limitations, and the correctness of the LCF is largely affected by the choice of the references used in the fitting procedure (Lombi & Susini, 2009; Tian et al., 2010). The references selected in this study principally include Zn association with organic acids which have been detected in the xylem sap of HP and NHP S. alfredii (Table 1), and a few other chemical forms of Zn. Therefore, Zn association with other chemicals, such as nicotianamine, cannot be ruled out completely.
Remobilization of metals by phloem transport has been studied less extensively than the xylem transport of metals in hyperaccumulator plants. The present work is, to our knowledge, the first to reveal Zn redistribution by phloem in hyperaccumulator species. Remobilization of Zn from old leaves to new growing leaves was investigated using stable isotope tracer technology, which has been applied successfully to investigate the phloem transport ability of Zn in rice plants by our research group (Wu et al., 2010). The results showed a 7.5-fold greater 68Zn transport from labeled mature leaves to the new growing leaves of HP relative to NHP S. alfredii (Fig. 4). Subsequent analysis by μ-XRF (Fig. 5), complemented by LA-ICPMS (Fig. 6), confirmed the enhanced ability of HP plants to remobilize Zn, and to preferentially distribute Zn to mesophyll cells surrounding phloem in the new leaves.
The results presented here demonstrate that plants of the Zn hyperaccumulator S. alfredii exhibit both enhanced xylem loading and xylem transport of Zn, as well as an enhanced ability for remobilization of the metal through phloem transport. Previously, Yang et al. (2006) reported that 65Zn transport is stimulated at the leaf cell plasma membrane in HP S. alfredii when exposed to a high Zn level, although no significant differences were observed at low Zn levels for HP and NHP S. alfredii. The results of Yang et al. (2006), however, only demonstrated stimulated leaf uptake of Zn into the leaf symplastic pool, and did not provide information on the possible enhancement of remobilization of Zn from the mature leaves to the new growing leaves. The processes involved in the phloem transport of Zn in HP S. alfredii need to be investigated in greater detail to elucidate the role of phloem remobilization of metals in hyperaccumulation. The high remobilization of 68Zn from mature to young leaves would not appear to be a favorable hyperaccumulation strategy, as it would enhance Zn levels in young developing tissues. It would be interesting to investigate whether the highly efficient remobilization of Zn results from a high requirement of Zn in the new growing tissues of the hyperaccumulators. Plants and plant cells of hyperaccumulators, including S. alfredii, have been suggested to have a higher requirement for the metals (Klein et al., 2008; Tian et al., 2009; Liu et al., 2010). Remobilization of mineral nutrients is important during the ontogenesis of a plant at periods of insufficient supply to the roots during vegetative growth (Marschner, 1995). The main sites (sources) for phloem loading of minerals are located in the mature stems and leaves as components of mineral nutrient supply to growth sinks (Marschner, 1995). The results of the present study confirm a high remobilization of Zn from the old mature leaves to the growth sinks in the hyperaccumulator S. alfredii, which probably results from a high metal requirement of the growing young leaves. It was not possible in the present study to quantify the role of vacuolar Zn remobilization in the hyperaccumulator. Given that vacuolar storage of metals is a general mechanism of metal sequestration in hyperaccumulators, further investigation of the potential re-usage of vacuolar Zn is warranted.
This work was supported by Projects from the National Natural Science Foundation of China (31000935, 21177107) and the Fundamental Research Funds for the Central Universities (2012FZA6008). Portions of this research were carried out at the Stanford Synchrotron Radiation Lightsource (SSRL), a Directorate of the Stanford Linear Accelerator Center (SLAC) National Accelerator Laboratory and an Office of Science User Facility operated for the US Department of Energy Office of Science by Stanford University. The SSRL Structural Molecular Biology Program is supported by the DOE Office of Biological and Environmental Research, and by the National Institutes of Health (NIH), National Institute of General Medical Sciences (NIGMS) (including P41GM103393) and the National Center for Research Resources (NCRR) (P41RR001209). The contents of this publication are solely the responsibility of the authors and do not necessarily represent the official views of NIGMS, NCRR or NIH. The authors sincerely thank all the staff of BL 7-3 and BL 2-3 at SSRL, particularly Serena DeBeer and Erik Nelson, for their support.