JXB Advance Access originally published online on March 14, 2005
Journal of Experimental Botany 2005 56(415):1343-1349; doi:10.1093/jxb/eri135
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RESEARCH PAPER |
Detection and quantification of ligands involved in nickel detoxification in a herbaceous Ni hyperaccumulator Stackhousia tryonii Bailey
1Primary Industries Research Centre, School of Biological and Environmental Sciences, Central Queensland University, Rockhampton, Queensland 4702, Australia
2School of Botany, The University of Melbourne, Melbourne, Victoria 3010, Australia
To whom correspondence should be addressed. Fax: +61 3 9349 4523. E-mail: ajmb{at}unimelb.edu.au
Received 22 November 2004; Accepted 13 February 2005
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Abstract |
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Field-collected, young plants of Ni hyperaccumulator Stackhousia tryonii, grown in a glasshouse for 20 weeks, were exposed to low- (available Ni concentration in the native serpentine soil, i.e. 60 µg g–1 dry soil) and high- (external application of 1000 ppm) Ni concentrations in the substrate. Nickel concentration in the freeze-dried leaf tissues increased from 3700 µg g–1 to 13 700 µg g–1 with soil Ni supplementation, of which >60% was extracted with dilute acid (0.025 M HCl). Nickel supplementation also elicited a 575%, 211%, and 37% increase in the final concentrations of oxalic, citric, and malic acids, respectively, in leaf tissues. Malic acid was the dominant organic acid, followed by citric and oxalic acids. The molar ratio of Ni to malic acid was 1.0, consistent with a role for malate as a ligand for Ni in hyperaccumulating plants, supporting detoxification/transport and storage of this heavy metal in S. tryonii. The total amino acid concentrations in the xylem sap did not change with Ni supplementation (21.7±3.7 mM and 17.9±5 mM, respectively, for low- and high-nickel-treated plants). Glutamine was the major amino acid in both the low- and high-Ni-treated plants. The concentration of glutamine decreased by >60%, with a corresponding increase in alanine, aspartic acid, and glutamic acid, on exposure to high Ni. A role of amino acids in Ni complexation and transport in S. tryonii is not immediately apparent.
Key words: Amino acids, metal hyperaccumulation, nickel (Ni), organic acids, Stackhousia tryonii Bailey
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Introduction |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
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Hyperaccumulation of heavy metals in plants is intriguing biologically, and is an extremely rare (exhibited by <0.2% of angiosperms; Baker et al., 2000
) phenomenon. Following uptake of heavy metals by the plant roots, translocation to the shoot and detoxification within the storage sites are two critical steps. This is achieved by chelation, transport, trafficking, and sequestration (Clemens, 2001; Clemens et al., 2002) by organo-ligands at a subcellular level. The potential ligands have been grouped into three major classes; oxygen donor ligands (e.g. carboxylates: malate, citrate, malonate, succinate, and oxalate), sulphur donor ligands (e.g. metallothioneins and phytochelatins), and nitrogen donor ligands (e.g. amino acids) (Baker et al., 2000). Complexation of metals with these ligands results in decreased free ion activity and thus reduced toxicity. Following detoxification, heavy metals are either compartmentalized in (epidermal) vacuoles (Brooks et al., 1981; Frey et al., 2000; Krämer et al., 2000; Persans et al., 2001), bound to the cell walls (Frey et al., 2000; Krämer et al., 2000), or localized in the apoplast (Vázquez et al., 1992).
It is clear that the ability of a ligand to lower the activity of the free metal ion is the key, and is highly pH-dependent. At acidic pH, organic acids are better chelators of Ni than amino acids, and as pH increases, the competition by amino acids increases until it exceeds the organic acids. It has been suggested (Homer et al., 1995) that hyperaccumulation of Ni cannot be accounted for by complexing with molecules of high molar mass (e.g. phytochelatins, metallothioneins) and that Ni appears to be associated with polar molecules of low molar mass (e.g. organic acids, amino acids). The involvement of heterocyclic nitrogen donors (amino acids) in the uptake of Ni2+ by plants has also been suggested (Still and Williams, 1980). Further, these nitrogen donor centres have higher stability constants for Ni than do carboxylates (Homer et al., 1995), and hence offer better transport of this metal within the system.
Organic acids
Nickel forms complexes with (in order of decreasing affinity): citric (Lee et al., 1977, 1978; Homer et al., 1991; Sagner et al., 1998; Bidwell, 2001; Schaumlöffel et al., 2003), malic (Brooks et al., 1981; Gabbrielli et al., 1991; Homer et al., 1991; Anderson et al., 1997), and malonic (Kersten et al., 1980; Brooks et al., 1981) acids.
Using a combination of advanced chromatographic and spectrometric techniques, Schaumlöffel et al. (2003) demonstrated that in the water extract of the latex of the Ni hyperaccumulating tree Niemeyera (Sebertia) acuminata, 99.4% of the Ni was sequestered by citrate and 0.3% by nicotinamine. Similarly, based on HPLC analysis of cellular sap and xylem fluids, it was suggested (Gabbrielli et al., 1997) that malate is produced in the roots, and then co-translocated with Ni to the shoots where it is preferentially localized in the leaf tissues. Salt et al. (2002) also suggested that Ni and Zn are transported to the shoot either as a metal–organic acid complex or as the hydrated cation. Once in the shoot, both Ni and Zn accumulate in the vacuole, where co-ordination by organic acids is favoured by the low pH (<6) value in the vacuole (Salt et al., 2002).
Amino acids
Reports on the role of amino acids in the hyperaccumulation of metals (including Ni) by plants are limited. Most studies failed to provide causative relationships between amino acid levels and metal hyperaccumulation (or tolerance). The production of particular amino acid(s) and their concentrations in hyperaccumulating plants, as a consequence of exposure to elevated levels of heavy metals (or hyperaccumulation), appears to be a species-specific phenomenon and is not necessarily present in all the metal hyperaccumulating plants. For example, Krämer et al. (1996) noted an increase in the concentration of histidine in the xylem sap of three Ni hyperaccumulators Alyssum lesbiacum, A. murale, and A. bertolonii, in proportion to external Ni application. Similarly, nickel application in the hydroponic solution elicited a large and proportional increase in free histidine in the leaves of Berkheya coddii (Harper et al., 1999). By contrast, in bulk leaf extracts of four Ni hyperaccumulators, Homer et al. (1995) noted only a slight change in the total amino acids over a wide range of leaf Ni concentrations. Farago and Mahmoud (1983) and Homer et al. (1995) failed to find any relationship between external Ni levels and amino acid content in the xylem fluids (or above-ground tissue) of hyperaccumulating plants, and concluded that the relationship between amino acid content and metal accumulation, if any, is obscure and enigmatic. Furthermore, while commenting on the need to minimize the risk of artefactual observations following tissue processing, Homer et al. (1995) emphasized the need to examine specific samples such as xylem sap for amino acid determinations, particularly for Ni hyperaccumulators.
Stackhousia tryonii Bailey is a rare, herbaceous Ni hyperaccumulator endemic to the serpentine soils of central Queensland (Australia). Bidwell (2001) studied organic acid concentrations in S. tryonii and reported citric acid to be the dominant ligand in the aqueous shoot extracts complexing only about one-third of the hyperaccumulated Ni (molar ratio of water-soluble Ni to citric acid=2.5:1; total Ni to citric acid=3.4:1). Bidwell (2001) concluded that further research was required to identify any additional compounds involved in the sequestration of the water-soluble Ni. Therefore, the present investigation was aimed at the identification and quantification of total (free) amino acids (xylem sap) and major organic acids (leaf tissue) as a consequence of the change in the external Ni concentration.
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Materials and methods |
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Plant material and growth conditions
Young plants of S. tryonii (having 2–3 branches), growing naturally in the serpentine soil near Eden Bann (23°05'72'' S, 105°16'41'' E; about 42 km north-west of Rockhampton, Queensland, Australia), were excavated. For excavation, 20 cm long pieces of PVC pipes (100 mm diameter) were vertically placed such that the plant was centred inside. PVC pipes were hammered into the soil (15 cm depth) by using a metal plate that rested on top of the pipe. PVC pipes with the intact plant in serpentine soil were withdrawn, brought to the glasshouse and placed in separate trays to contain drained out irrigation water or Ni solution. Each PVC pipe contained one plant. No fertilizers were added to the collected soils. The clusters of stems originating from a common base were carefully cut such that the root system was left intact (undisturbed) in the soil (within the PVC pipes) as a perennating structure. The temperature inside the glasshouse, during the growth period, varied between 12 °C (night) and 25 °C (day), and the relative humidity was approximately 60%.
Treatments
The soil was maintained at 60% field capacity (irrigated with r/o water) for 6–8 weeks, until plants produced 3–4 new stems (branches) from the perennating structure. Plants were randomly chosen for exposure to two levels of Ni treatments. ‘Low Ni’ plants received no external application of nickel. The ‘high Ni’ plants received 1000 ml of 1000 µg ml–1 nickel (as NiSO4.6H2O) supplied evenly over a 20 week growth period, giving a final (available) soil Ni concentration at 400 µg g–1 dry soil. Each treatment was replicated three times.
Tissue preparation
Plants were harvested after 20 weeks of growth. Shoots were cut at the shoot–root junction using a sharp razor blade, briefly immersed in r/o water, blotted-dry, and immediately plunged in liquid nitrogen. The frozen samples were freeze-dried at –98 °C for 100 h in a freeze-drier. Following freeze-drying, leaves were separated from stems and powdered separately in a ball mill.
Preparation of tissue extracts and HPLC analysis of organic acids
Plant extracts were obtained following the method of Tolrà et al. (1996). Briefly, to extract and dissociate the Ni–organic acid complexes, powdered leaf subsamples (100–200 mg) were mixed with 1.5 ml aliquots of 0.025 M HCl in sterile Eppendorf vials, sonicated for 10 min, and vortexed for 1 min. The extraction was carried out at ambient conditions and the procedure was repeated three times. The three extracts from each subsample were pooled and centrifuged at 13 000 g for 15 min. The final volume of the extracts was made to 5 ml with 0.025 M HCl and analysed individually for various organic acids by ion exclusion High Performance Liquid Chromatography (HPLC; Tolrà et al., 1996).
The HPLC consisted of a HPLC pump (Waters Corporation, MA, USA), an auto-injector (Shimadzu Corp., Kyoto, Japan) and a tunable absorbance detector. The system was interfaced to a Delta 5.0 Chromatography data system (Digital Solutions, Queensland, Australia). For all separations, an Aminex HPX-87H column (300 x 7.8 mm internal diameter) (Bio-Rad, Hercules, CA, USA), protected by a Bio-Rad cation H guard cartridge, was used with 0.003 N sulphuric acid as the mobile phase. Duplicate 10 µl injections were made of each standard and sample. The operating conditions of HPLC were as follows: flow rate 0.6 ml min–1, temperature 31 °C, elution time 26 min, and wavelength 210 nm. Organic acids in the plant extracts were identified (and quantified) by comparison of component retention times in standard solutions and peak integration to fit into standard curves for quantification (Tolrà et al., 1996).
All samples (analysed in duplicate for three replicates for both low- and high-Ni treatments) revealed early elution of high-UV-absorbing compounds, which distort the baseline (data not shown). Oxalic acid was therefore difficult to resolve. As a result, quantitation of oxalic acid is only approximate. Malic acid showed a slight concentration-dependent shift in retention time, and therefore a sample was spiked with malic acid standard and this verified the identification of the malic acid.
Xylem sap collection and HPLC analysis of amino acids
Xylem sap was collected using a pressure vessel (model 3000, Soil Moisture Equipment Corp., CA, USA). This method of xylem sap collection is different from ‘bleeding sap’ method. Briefly, branches (2–4 shoots per plant) were carefully severed at the shoot–root junction and immediately enclosed in a pressure vessel. The pressure was increased (0.1 MPa min–1) slightly above the balancing pressure and then maintained for 8–10 min. The first 10–20 µl of sap was discarded and the cut surface wiped with moist absorbent tissue to reduce contaminants. Thereafter, exuded xylem sap (350–600 µl in total from 2–4 branches) was collected by micropipette in sterile 1.5 ml Eppendorf vials (on ice), and stored frozen until analyses.
Free amino acids in the xylem sap sample were analysed using a Waters AccQTag amino acid analysis system. The amino acids in the xylem sap were separated by a sensitive precolumn derivatization technique that is used with reverse phase high performance liquid chromatography (RP-HPLC) and quantified by fluorescence or UV detection. Before separation, amino acids were derivatized with 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate (AQC; Cohen, 2001) which were then transferred to the HPLC auto-sampler for analysis.
The HPLC consisted of (in series): a separation module (Waters Corporation, MA, USA), a fluorescence detector, and a dual 
absorbance detector. The control and analysis software was Waters Empower Pro. For all separations, a Waters AccQTag column (15 cmx3.9 mm internal diameter) was used. The column and auto-sampler temperatures were maintained at 37 °C and 10 °C, respectively. The sample injection volume and flow rate were 5 µl and 1 ml min–1, respectively. The fluorescence detector wavelengths were 250 nm (excitation) and 395 nm (emission). The wavelength of the UV/Vis detector was set at 254 nm.
Nickel analysis
For chemical analysis, plant extracts (as described earlier; 0.025 M HCl; 3 ml) were mixed with 70% (v/v) nitric acid and incubated in hot-water bath at 90 °C for 4 h. Digests were cooled to room temperature and 1–2 ml hydrogen peroxide was added. Digests were further incubated at 90 °C until clear and analysed for Ni using an inductively coupled plasma-optical emission spectrophotometer (Varian Vista AX; Varian Inc., Australia). Appropriate blanks and standard tissue samples were run to ensure analytical integrity.
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Results |
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Nickel hyperaccumulation
External Ni application elicited a dramatic increment in Ni concentration in the leaf tissues (Table 1). The leaves of high-Ni-treated plants possessed 3.7 times as much Ni (at 13 700 µg g–1 dry weight) compared with low-Ni-treated plants (at 3700 µg g–1 dry weight). Similarly, dilute acid solution (0.025 M HCl) extracted four times as much Ni from the leaves of high-Ni-treated plants (at 9200 µg g–1 dry weight) compared with low-Ni-treated plants (at 2250 µg g–1 dry weight). The dilute acid solution extracted 61% and 68% Ni from the leaves of low- and high-Ni-treated plants, respectively.
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Organic acids
The leaf extracts subjected to HPLC analysis were tested for five predominant organic acids: oxalic, citric, malonic, malic, and succinic. However, of these five, only oxalic, citric, and malic acids were detectable in the samples (data not shown). The most abundant organic acid in the leaves of low- and high-Ni-treated plants was malic acid (21 700 and 29 700 µg g–1, respectively), followed by citric (2650 and 8250 µg g–1, respectively) and oxalic (110 and 750 µg g–1, respectively) acids (Table 1). Treatment of plants with a high external Ni (1000 ppm) application elicited elevated concentrations of all the organic acids tested in the leaf extracts. This increment in the concentration of oxalic, citric, and malic acids was about 575%, 211%, and 37%, respectively. The concentrations of all organic acids varied greatly across replicates within each of the two treatments, as shown by the high values of standard error (Table 1).
The molar ratios of both the bulk (total) and dilute acid-extractable Ni to organic acids followed the order: oxalic>citric>malic.
Amino acids
A total of 19 free amino acids were detected in the xylem sap (Table 2). In the xylem sap of low-Ni-treated plants, glutamine was the dominant amino acid, with concentrations reaching up to 11 mM, and accounting for half of the total amino acid concentrations in the xylem sap (Table 2). Glutamic acid, alanine, and aspartic acid were also present but their concentrations were only 2.8, 1.2 and 1.1 mM, respectively. All other amino acids were present in very low concentrations (<1.0 mM).
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In the xylem sap of high-Ni-treated plants glutamine, glutamic acid, alanine, and aspartic acid were the major amino acids at 4.1, 3.4, 2.9, and 1.5 mM, respectively. Glutamine accounted for nearly one-quarter of the total amino acid concentration in the xylem sap. The individual concentrations of all other amino acids were below 1.0 mM (Table 2).
The total concentration of free amino acids in the xylem sap was similar in low- (21.7±3.7 mM) and high-Ni-treated plants (17.9±5 mM).
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Discussion |
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Tissue analysis involves the homogenization of many cellular compartments. By contrast, xylem sap represents a single, specific compartment. Unfortunately, small volumes of xylem sap were extracted, permitting the analysis of amino acids, but not of Ni and organic acids. Future studies should consider mechanisms to allow the analysis of smaller volumes, or to allow the extraction of greater volumes of xylem sap (e.g. bulking of samples from replicates).
Leaf nickel concentration
The nickel concentrations measured in the leaf tissues of S. tryonii are in line with those reported earlier for this species (Batianoff et al., 1990; Bidwell, 2001). Plants accumulated 3.5 times more Ni on exposure to high available Ni in the substrate. Between 60% and 70% of the bulk (total) nickel in the leaf tissue was dilute acid (0.025 M HCl) soluble. Bidwell (2001) extracted up to 72% water-soluble nickel from the whole shoot tissues of S. tryonii. For freeze-dried stem tissues, similar results were also obtained (64% and 76% for low- and high-Ni-treated plants; data not presented) using dilute acid as the extractant. A water wash of the thin (c. 20 µm) hand-sectioned specimens of leaf and stem extracted between 73% and 92% of Ni (as determined using micro-PIXE technique; Bhatia et al., 2004). The lower extraction of Ni reported here was possibly due to inadequate extraction of Ni from the freeze-dried leaves or the age of the specimens used in the two studies. Mature plants possess comparatively higher cellulose/pectic material, and are therefore expected to hold larger amounts of Ni than the younger ones.
Presumably, the remaining Ni was associated with the cell wall, as reported for another Ni hyperaccumulator Hybanthus floribundus (Farago and Mahmoud, 1983). Similarly, up to 68% of the total Ni in the hyperaccumulator Thlaspi goesingense was associated with the cell wall material (Krämer et al., 2000). Higher concentrations of acid are expected to displace substantial amounts of Ni from negatively charged sites on the cell wall.
Organic acids (leaf tissue)
Levels of oxalic, citric, and malic acids increased substantially when the plants were exposed to higher external Ni concentrations. However, only malic acid was present in concentrations equivalent to that of Ni in the plant. Malate has previously been reported to sequester Ni in several hyperaccumulating plants. For example, Anderson et al. (1997) reported that in the leaves of the Ni hyperaccumulator Berkheya coddii, malate co-ordinated with Ni in a 1:1 molar ratio. Homer et al. (1991) also noted a mole ratio of 1:0.4:1 (Ni:citric acid:malic acid) in a purified extract of the hyperaccumulator Dichapetalum gelonioides ssp. tuberculatum, and demonstrated that the Ni–citrate complex was more stable, but that the amount of citrate was insufficient to complex all the Ni.
The molar ratios of Ni to citric acid (between 3:1 and 6:1) in the bulk (leaf) tissues and dilute acid extract were in general agreement with that reported by Bidwell (2001) for shoot tissues (between 2.5:1 and 4.8:1). Bidwell (2001) used HPLC and GC-MS techniques on Ni-rich fractions collected from the G-10 column, and detected oxalic, citric, malic, and malonic acids in the stem extracts (0.025 M HCl). However, only citric acid was quantified, as this was apparently the dominant organic acid – sequestering about one-third of the total Ni and 50% of the water-extractable Ni. By contrast, in the current study, malic acid was the predominant organic acid in the bulk leaf tissue. This difference may relate to the type of tissue used (bulk leaf versus shoot), or may represent a misinterpretation (Bidwell did not attempt to quantify malic acid). In addition, Reeves (1992) suggested that malate may be partially destroyed and reconstituted by the dilution and re-concentration processes during separation.
Bidwell (2001) concluded that further research was required to detect additional compounds involved in the sequestration of Ni in S. tryonii. The present study supports the assertion that citric acid may have significance to the complexation of Ni (molar ratio equivalent to about 20% of Ni); however, malic acid was recorded in higher concentration, although Reeves (1992) has noted that the equilibrium constant for the binding of Ni to malate was 150 times smaller than citrate.
Amino acids (xylem sap)
In the current study, total amino acid concentration decreased with Ni treatment. However, the concentration of solutes in the xylem sap is dependent on transpiration rates and such concentration values should not be used to compare treatments. A preferable index is the percentage composition of xylem sap.
Glutamine was the dominant amino acid in the xylem sap of low-Ni-treated plants, with concentrations of glutamic acid, aspartic acid, and alanine increased at high external Ni concentration. In light of this, presumably the exposure of S. tryonii plants to elevated Ni concentrations resulted in the induction of glutamate synthase (GOGAT) and amino transferase, both of which assist in the conversion of glutamine through glutamate into amino acids. Nickel also appears to have induced a substantial shift in the operative pathways of ammonia assimilation, and the induction of GOGAT activity under Ni stress may provide the glutamate required for enhancing the synthesis of amino acids, such as glutamic acid, alanine, and aspartic acid. El-Shintinawy and El-Ansary (2000) studied amino acid metabolism in soybean, and reported that high Ni levels in the xylem sap seems to account only for the decrease in alanine aminotransferase (ALT) and aspartate aminotransferase (AST), both of which contribute to the accumulation of alanine and aspartic acid, respectively. However, further studies are warranted to confirm this in S. tryonii.
The stability constant for glutamic acid is low. It is possible that other amino acids confer a higher Ni transport capacity to the xylem sap. These changes may increase the Ni carrying capacity of the xylem sap, with Homer et al. (1995) reporting higher stability constants for Ni and aspartic acid and alanine than other amino acids (glutamine?). It would be useful to establish the stability constant for glutamine.
A proportional increase in histidine concentrations in the xylem sap to external Ni application accounts for the detoxification and translocation of Ni in three Ni hyperaccumulators Alyssum lesbiacum, A. murale, and A. bertolonii (Krämer et al., 1996). The stability constant for histidine is high but histidine was not produced in sufficient quantities in the xylem sap of S. tryonii, even with the higher external application of Ni. Clearly, S. tryonii behaves differently than these Ni hyperaccumulators, indicating that histidine is not involved in Ni detoxification or translocation in this species.
Cysteine was not detected in the xylem sap of S. tryonii. It is unlikely that phytochelatins are involved in Ni detoxification/hyperaccumulation of this metal in this species. Moreover, recently, it has been demonstrated (Vatamaniuk et al., 2000; Ebbs et al., 2002; Schat et al., 2002) that phytochelatins are not involved in metal tolerance or accumulation in hyperaccumulating plants (especially Ni and Zn hyperaccumulators). For example, Sagner et al. (1998) reported that the phytochelatin synthase [enzyme responsible for the synthesis of (
-glu-cys)n-gly] is not activated by Ni in hyperaccumulators. By contrast, Mesjasz-Przybylowicz et al. (2003) noted elevated concentrations of phytochelatins in the leaves of Ni hyperaccumulator Berkheya coddii. To resolve the issue of whether or not phytochelatins are involved in Ni detoxification in S. tryonii, further research is currently underway in this laboratory to use synchrotron-based (and other biochemical) techniques, which will provide insight on in planta co-ordination environment of Ni in the tissues.
Xylem sap organic acid levels may also have increased with Ni treatment. Since the regulation of nitrogen metabolism and amino acid metabolism are interconnected, it is difficult to envisage a precise physiological explanation to describe changes in amino acid levels in the xylem sap of S. tryonii, following exposure to high-Ni treatment. To elucidate precisely what role organic acids play in the tolerance, and detoxification, translocation, and storage of hyperaccumulated Ni in S. tryonii, it would have been useful to measure Ni and organic acid concentrations in the xylem sap. Future work should consider the monitoring of organic acid levels in the xylem sap in response to external Ni treatment.
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Acknowledgements |
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The merit scholarship (UPRA) to NPB by the Central Queensland University is gratefully acknowledged. We thank Professor JAC Smith (Oxford University) for review of the manuscript and Augustine Doronila (University of Melbourne) for help in chemical analysis (ICP-OES) of plant samples. The assistance of Bernie McInerney and Chris Clarke (Australian Proteome Analysis Facility, Macquarie University, Sydney) and Malcolm Nobel (University of New South Wales, Sydney) for amino acid and organic acid analyses is thankfully acknowledged. Thanks are also due to the Queensland Parks and Wildlife Services (Central Region) for granting a scientific permit to collect S. tryonii material from its natural habitat.
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Footnotes |
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* Present address: Environment Division, Australian Nuclear Science and Technology Organisation (ANSTO), PMB 1 Lucas Heights, Menai, NSW 2234, Australia.
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References |
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TopAbstractIntroductionMaterials and methodsResultsDiscussionReferences |
|---|
Anderson TR, Howes AW, Slatter K, Dutton MF. 1997. Studies on the nickel hyperaccumulator, Berkheya coddii. In: Jaffré T, Reeves RD, Becquer T, eds. The ecology of ultramafic and metalliferous areas. Proceedings of the Second International Conference on Serpentine Ecology, Nouméa. Nouméa, New Caledonia: ORSTOM, 261–266.
Baker AJM, McGrath SP, Reeves RD, Smith JAC. 2000. Metal hyperaccumulator plants: a review of the ecology and physiology of a biological resource for phytoremediation of metal-polluted soils. In: Terry N, Bañuelos G, eds. Phytoremediation of contaminated soil and water. Boca Raton, Florida, USA: Lewis Publishers, 85–107.
Batianoff GN, Reeves RD, Specht RL. 1990. Stackhousia tryonii Bailey: a nickel-accumulating serpentine-endemic species of Central Queensland. Australian Journal of Botany 38, 121–130.[CrossRef]
Bhatia NP, Walsh KB, Orlic I, Siegele R, Ashwath N, Baker AJM. 2004. Studies on spatial distribution of nickel in leaves and stems of the metal hyperaccumulator Stackhousia tryonii Bailey using nuclear microprobe (micro-PIXE) and EDXS techniques. Functional Plant Biology 31, 1061–1074.[CrossRef]
Bidwell SD. 2001. Hyperaccumulation of metals in Australian native plants. PhD thesis, The University of Melbourne, Australia.
Brooks RR, Shaw S, Asensi Marfil A. 1981. The chemical form and physiological function of nickel in some Iberian Alyssum species. Physiologia Plantarum 51, 167–170.[CrossRef]
Clemens S. 2001. Molecular mechanisms of plant metal tolerance and homeostasis. Planta 212, 475–486.[CrossRef][Web of Science][Medline]
Clemens S, Palmgren MG, Krämer U. 2002. A long way ahead: understanding and engineering plant metal accumulation. Trends in Plant Science 7, 309–315.[CrossRef][Web of Science][Medline]
Cohen SA. 2001. Amino acid analysis using precolumn derivatisation with 6-aminoquinolyl-N-hydroxysuccinimidyl carbamate. In: Cooper C, Packer N, Williams K, eds. Methods in molecular biology, Vol. 159. Totowa, New Jersey, USA: Humana Press, 39–47.
Ebbs S, Lau I, Ahner B, Kochian L. 2002. Phytochelatin synthesis is not responsible for Cd tolerance in the Zn/Cd hyperaccumulator Thlaspi caerulescens (J.&C. Presl). Planta 214, 635–640.[CrossRef][Web of Science][Medline]
El-Shintinawy F, El-Ansary A. 2000. Differential effect of Cd2+ and Ni2+ on amino acid metabolism in soybean seedlings. Biologia Plantarum 43, 79–84.
Farago ME, Mahmoud IEDAW. 1983. Plants that accumulate metals. VI. Further studies of an Australian nickel accumulating plant. Minerals and the Environment 5, 113–121.
Frey B, Keller C, Zierold K, Schulin R. 2000. Distribution of Zn in functionally different leaf epidermal cells of the hyperaccumulator Thlaspi caerulescens. Plant, Cell and Environment 23, 675–687.[CrossRef]
Gabbrielli R, Gremigni P, Bonzi Morassi L, Pandolfini T, Medeghini P. 1997. Some aspects of Ni tolerance in Alyssum bertolonii Desv.: strategies of metal distribution and accumulation. In: Jaffré T, Reeves RD, Becquer T, eds. The ecology of ultramafic and metalliferous areas. Proceedings of the Second International Conference on Serpentine Ecology, Nouméa. Nouméa, New Caledonia: ORSTOM, 225–227.
Gabbrielli R, Mattioni C, Vergnano O. 1991. Accumulation mechanisms and heavy metal tolerance of a nickel hyperaccumulator. Journal of Plant Nutrition 14, 1067–1080.
Harper FA, Baker AJM, Balkwill K, Smith JAC. 1999. Nickel uptake, translocation and hyperaccumulation in Berkheya coddii. In: Abstracts of the Third International Conference on Serpentine Ecology, Kruger National Park, South Africa.
Homer FA, Reeves RD, Brooks RR, Baker AJM. 1991. Characterisation of the nickel-rich extract from the nickel hyperaccumulator Dichapetalum gelonioides. Phytochemistry 30, 2141–2145.[CrossRef]
Homer FA, Reeves RD, Brooks RR. 1995. The possible involvement of amino acids in nickel chelation in some nickel-accumulating plants. Current Topics in Phytochemistry 14, 31–37.
Kersten WJ, Brooks RR, Reeves RD, Jaffré T. 1980. Nature of nickel complexes in Psychotria douarrei and other nickel-accumulating plants. Phytochemistry 19, 1963–1965.[CrossRef]
Krämer U, Cotter-Howells JD, Charnock JM, Baker AJM, Smith JAC. 1996. Free histidine as a metal chelator in plants that accumulate nickel. Nature 379, 635–638.[CrossRef]
Krämer U, Pickering IJ, Prince RC, Raskin I, Salt DE. 2000. Subcellular localization and speciation of nickel in hyperaccumulator and non-accumulator Thlaspi species. Plant Physiology 122, 1343–1353.
Lee J, Reeves R, Brooks R, Jaffré T. 1977. Isolation and identification of a citrato-complex of nickel from nickel-accumulating plants. Phytochemistry 16, 1503–1505.[CrossRef]
Lee J, Reeves R, Brooks R, Jaffré T. 1978. The relation between nickel and citric acid in some nickel-accumulating plants. Phytochemistry 17, 1033–1035.[CrossRef]
Mesjasz-Przybylowicz JM, Stroi
ski A, Chadzinikolau T. 2003. Phytochelatins in Berkheya coddii nickel hyperaccumulating plant from South Africa. In: Abstracts of the Fourth International Conference on Serpentine Ecology, Havana, Cuba.
Persans MW, Nieman K, Salt DE. 2001. Functional activity and role of cation-efflux family members in Ni hyperaccumulation in Thlaspi goesingense. Proceedings of the National Academy of Sciences, USA 98, 9995–10000.
Reeves RD. 1992. Hyperaccumulation of nickel by serpentine plants. In: Baker AJM, Proctor J, Reeves RD, eds. The vegetation of ultramafic (serpentine) soils. Proceedings of the First International Conference on Serpentine Ecology. Andover, UK: Intercept Ltd., 253–277.
Sagner S, Kneer R, Wanner G, Cosson JP, Deus-Neumann B, Zenk MH. 1998. Hyperaccumulation, complexation and distribution of nickel in Sebertia acuminata. Phytochemistry 47, 339–347.[CrossRef][Web of Science][Medline]
Salt DE, Prince RC, Pickering IJ. 2002. Chemical speciation of accumulated metals in plants: evidence from X-ray absorption spectroscopy. Microchemical Journal 71, 255–259.[CrossRef]
Schat H, Llugany M, Vooijs R, Hartley-Whitaker J, Bleeker PM. 2002. The role of phytochelatins in constitutive and adaptive heavy metal tolerances in hyperaccumulator and non-hyperaccumulator metallophytes. Journal of Experimental Botany 53, 2381–2392.
Schaumlöffel D, Ouerdane L, Bouyssiere B, Lobinski R. 2003. Speciation analysis of nickel in the latex of a hyperaccumulating tree Sebertia acuminata by HPLC and CZE with ICP MS and electrospray MS-MS detection. Journal of Analytical Atomic Spectrometry 18, 120–127.[CrossRef]
Still ER, Williams RJP. 1980. Potential methods for selective accumulation of nickel (II) ion by plants. Journal of Inorganic Biochemistry 13, 35–40.
Tolrà RP, Poschenrieder C, Barceló J. 1996. Zinc hyperaccumulation in Thlaspi caerulescens. 2. Influence on organic acids. Journal of Plant Nutrition 19, 1541–1550.
Vázquez MD, Barceló J, Poschenrieder C, Mádicó J, Hatton P, Baker AJM, Cope GH. 1992. Localization of zinc and cadmium in Thlaspi caerulescens (Brassicaceae), a metallophyte that can hyperaccumulate both metals. Journal of Plant Physiology 140, 350–355.
Vatamaniuk OK, Mari S, Lu Y-P, Rea PA. 2000. Mechanism of heavy metal ion activation of phytochelatin (PC) synthase: Blocked thiols are sufficient for PC synthase-catalyzed transpeptidation of glutathione and related thiol peptides. Journal of Biological Chemistry 275, 31451–31459.
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