JXB Advance Access originally published online on February 5, 2007
Journal of Experimental Botany 2007 58(6):1333-1338; doi:10.1093/jxb/erl300
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© 2007 The Author(s).
This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/by-nc/2.0/uk/) which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.This paper is available online free of all access charges (see http://jxb.oxfordjournals.org/open_access.html for further details)
RESEARCH PAPER |
Can arsenic–phytochelatin complex formation be used as an indicator for toxicity in Helianthus annuus?
1Department of Chemistry, University of Aberdeen, Meston Building, Meston Walk, Aberdeen AB24 3UE, UK
2School of Biological Sciences, University of Aberdeen, Cruickshank Building, St Machar Drive, Aberdeen AB24 3UU, UK
* To whom correspondence should be addressed. E-mail: j.feldmann{at}abdn.ac.uk
Received 25 September 2006; Revised 11 December 2006 Accepted 15 December 2006
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Abstract |
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The formation of arsenic–phytochelatin (As–PC) complexes is thought to be part of the plant detoxification strategy for arsenic. This work examines (i) the arsenic (As) concentration-dependent formation of As–PC complex formation and (ii) redistribution and metabolism of As after arrested As uptake in Helianthus annuus. HPLC with parallel ICP-MS/ES-MS detection was used to identify and quantify the species present in plant extracts exposed to arsenate (As(V)) (between 0 and 66.7 µmol As l–1 for 24 h). At As concentrations below the EC50 value for root growth (22 µmol As l–1) As uptake is exponential, but it is reduced at concentrations above. Translocation between root and shoot seemed to be limited to the uptake phase of arsenic. No redistribution of As between root and shoot was observed after arresting As exposure. The formation of As–PC complexes was concentration-dependent. The amount and number of As–PC complexes increased exponentially with concentration up to 13.7 µmol As l–1. As(III)–PC3 and GS–As(III)–PC2 complexes were the dominant species in all samples. The ratio of PC-bound As to unbound As increased up to 1.3 µmol As l–1 and decreased at higher concentrations. Methylation of inorganic As was only a minor pathway in H. annuus with about 1% As methylated over a 32 d period. The concentration dependence of As–PC complex formation, amount of unbound reduced and oxidized PC2, and the relative uptake rate showed that As starts to influence the cellular metabolism of H. annuus negatively at As concentrations well below the EC50 value determined by more traditional means. Generally, As–PC complexes and PC-synthesis rate seem to be the more sensitive parameters to be studied when As toxicity values are to be estimated.
Key words: Arsenic, arsenic–phytochelatin complex, arsenic speciation, ES-MS, Helianthus annuus, ICP-MS, methylation, phytochelatin
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Introduction |
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Although it has been suspected for some time that phytochelatins (PCs) provide a vital role in plant defence against arsenic and other potentially toxic elements with high affinity for the PCs cysteine S-H functional groups, there is little direct evidence for the actual role of As–PC complexation in planta, the significance of As–PC complexation has not been established in planta (Rauser, 1995; Sneller et al., 1999; Cobbett, 2000; Pickering et al., 2000; Hartley-Whitaker et al., 2001; Meharg and Hartley-Whitaker, 2002; Bleeker et al., 2003, 2006). Raab et al. (2004, 2005) have developed analytical approaches, HPLC-ICP-MS/ES-MS, to identify plant-extracted As–PC complexes, showing for the first time the nature and diversity of these complexes. In particular, detailed time-course experiments with sunflower (Helianthus annuus) revealed the sequence of phytochelatin (PC) complex formation, the general diversity of As–PC complexes in cells, de novo methylation of inorganic arsenic and subsequent PC-complexation, and lack of involvement of PCs in xylem transport of arsenic (Raab et al., 2005). It has also been shown recently that As–PC complexes may have a vital role in translocating As to vacuoles, probably through tonoplast ABC-type transporters (Bleeker et al., 2006).
It is crucial when studying the interaction of a toxicant with a physiological process to understand the toxicants dose response. Arsenic species directly interact with cellular components to disrupt physiological processes required for cell function, such as binding to protein S-H groups and arsenate substituting for phosphate in ATP synthesis, given that arsenate is rapidly converted to arsenite in plant cells (Meharg and Hartley-Whitaker 2002), and the redox-cycling between As(III) and As(V) species causes oxidative stress.
As–PC synthesis places a strong demand on plant S stores (McMahon and Anderson, 1998). The general stress caused by arsenic exposure will have inevitable consequences for specific biochemical pathways. Metabolism of inorganic arsenic through methylation may be an important detoxification mechanism, and it is also known that PC-methylated arsenic species complexes can form (Raab et al., 2005).
This paper stresses how arsenate concentration affects PC complex formation and arsenic methylation in sunflowers using HPLC-ICP-MS/ES-MS, and how this speciation relates to As toxicity.
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Materials and methods |
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Plants and growth conditions
H. annuus plants (cultivar Giant Yellow) were cultivated on Perlit®-substrate in a greenhouse at 12 h light/dark cycle at 18/16 °C, watered and fertilized with Hoagland solution (1.25 mM KNO3, 1.50 mM Ca(NO3)2, 0.75 mM MgSO4, 0.50 mM KH2PO4, 50 µM KCl, 50 µM H3BO3, 10 µM MnSO4, 2.0 µM ZnSO4, 1.5 µM CuSO4, 0.5 µM Na2MoO4.2H20, 0.1 mM Na2O3Si, and 72 µM Fe-EDTA pH 6.0) every 2 d. After 6 weeks plants were grown for a further 7 d without fertilization before exposure to As (100 ml of 0–66.7 µmol l–1 As(V) in water).
All plants were harvested 24 h after As exposure. The total fresh-weight (FW) of the plants used was not significantly different (P >0.3 in all cases, Students t test).
For the determination of the long-term As distribution, plants were grown for 2 weeks in Perlit® and fertilized regularly with Hoagland solution. Between weeks 2 and 3 the plants were not fertilized and at the end of week 3 they were exposed to 1.3 µmol l–1 As(V) in water (100 ml) for 24 h and then repotted in fresh Perlit®. Plants were harvested 1, 2, 4, 8, 16, or 32 d later. During that time plants were fertilized every 3 d. This experiment was also intended to study methylation of As by H. annuus.
Plants grown for the short-term toxicity test were germinated on filter-paper and exposure to varying arsenate (0–100 µmol l–1 As(V)) concentrations started after 3 d using hydroponic culture conditions as described in more detail in Abedin and Meharg (2002). Throughout, the plants (n=9 for each concentration) were exposed for 7 d to arsenate, after which root length and root weight were recorded.
Sample preparation
The plants were prepared for analysis immediately following harvest. Root and shoot were separated and, after washing of the root [10 min in 10 mM potassium hydrogen phosphate, 10 min in distilled water, similar to Meharg and MacNair (1992)], ground using liquid nitrogen, and subsampled. A subsample of the material was immediately extracted using 1% formic acid (90 min at 1 °C, then filtered) for the determination of As–PC complexes. Samples intended for the determination of methylated As compounds were extracted in the same way, and more details are given in Raab et al. (2004).
Other subsamples were digested using nitric acid/hydrogen peroxide (digestion on a hot plate until clear digest was obtained) for the determination of total arsenic.
Arsenic speciation analysis
For the separation, identification, and quantification of the As species present in extracts, a combination of HPLC-ICP-MS/ES-MS (C18, Spherisorb S5 ODS2 250 mmx4.6 mm, Waters USA) was used within a few hours after extraction. A gradient of 1% formic acid in water and methanol was used for the separation and the eluent-flow was split after the column (1 part into the ICP-MS (Agilent 7500c, Agilent USA) and 4 parts into the ES-MS (Agilent 1100 MSD, Agilent USA)). The ES-MS was used in the positive single ion-monitoring mode with a capillary voltage of 4000 V and a fragmentor voltage of 100 V.
Anion-exchange chromatography (PRP X-100 Hamilton, 150 mmx4.6 mm, Hamilton USA) with 20 mM ammonium-phosphate buffer pH 6.0 as eluent and HPLC-ICP-MS was used to determine dimethylarsinic acid (DMA(V)), monomethylarsonic acid (MA(V)), and inorganic As. Standard-addition was used to verify the identity of DMA(V) and MA(V). External calibration with DMA(V) was used in both cases for quantification.
Total As in digests and extracts was measured by ICP-MS (Agilent 7500c, Agilent, USA) using external calibration for the determination of extraction efficiency and chromatographic recovery.
Data analysis
All data are presented on a plant fresh weight basis. Quantification was done using the ICP-MS signal on m/z 75, for total As external calibration was used and for speciation DMA(V) (peak areas versus concentration), the determination limit (10 
) for total As was 0.0001 µmol g–1 plant FW and for speciation 0.005 µmol g–1 plant FW. For total As IAEA 140 (Fucus spp) was used as reference material with a certified As concentration of 44.3±4.2 mg kg–1 (measured As concentration 40.7±1.3 mg kg–1, n=20). No reference material exists for verification of the quantification of As–PC complexes. These were verified using extraction efficiency and chromatographic recovery, both of which were 92±9% (n=42).
Root tolerance index (RTI) was used for the expression of As(V) toxicity. It was calculated by dividing the length of the As treated roots by that of the control plants multiplied by 100, the same calculation was used utilizing root weight. Root length growth has been used by various authors (Abedin and Meharg, 2002) to determine the toxicity of arsenate. All experiments were done in triplicate and statistical analysis was done by Students t test (P <0.05 significance level).
To estimate the As concentration dependence of GSH, GSSG, and reduced and oxidized PC2, peak areas measured by ES-MS were used. The peak areas of the GS–As(III)–PC2 and the As(III)–PC3 complex as measured by ES-MS were correlated with the concentration of these complexes as determined by the arsenic signal of the ICP-MS measurements in all samples. The slope and intercept of these calculations were then used to calculate GSH, GSSG, and unbound reduced and oxidized PC2 concentration changes depending on As concentration. Free reduced or oxidized PC3 was below the detection limit of ES-MS.
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Results |
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Dose-dependent arsenic uptake over 24 h
Generally, roots contained between 50% and 85% of the total amount of As present in the plant, the rest was relative equally distributed between stem and leaf. The relative uptake per whole plant was highest (59±18% of available As) during exposure to 0.13 µmol As l–1. Plants exposed to 66.7 µmol As l–1 took up 27±7% of available As (Fig. 1). A linear relationship between exposure and concentration in the plant was found for leaf and stem; the relationship for root was best described as an increase to maximum. Percentage uptake of As for the whole plant was best described by a combination of exponential rise followed by a reduction with the reduction starting around 13.3 µmol l–1 As exposure. Statistical analysis (Student's t test, significant at P <0.05) of the data showed that the As concentration in roots differed significantly with all As concentration, whereas no significant difference in total As concentration (Fig. 1) in leaf and stem of plants exposed to less than 1.3 µmol l–1 was detectable.
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Long-term distribution of total As after one-time exposure to As(V)
The long-term distribution experiment showed that H. annuus did not excrete As back into its rooting environment. There was no decrease in total As per plant over time (Fig. 2). After initial As uptake and translocation within the plant, there was no redistribution from root to leaf or vice versa (Fig. 2). The increase in As on the first day resulted from the fact that the roots of all exposed plants were not washed with phosphate-solution to remove externally attached As after exposure, instead they were washed during final sample preparation.
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As–PC complexes in dependence of varying As(V) concentrations
The root system was the predominant site of As–PC complex formation. An example of a root extract separation is shown in Fig. 3. As(III)–PC3 was the main complex formed after 24 h exposure to As(V), followed by GS–As(III)–PC2 at all tested concentrations except at 66.7 µmol As l–1, where GS–As(III)–PC2 concentration was higher. At As concentrations higher than 1.3 µmol As l–1 As(III)–(PC2)2 complexes were detectable in two different isomers. The MA(III)–PC2 complex was detectable in root extract of plants exposed to 6.7 µmol As l–1 or more. A number of other species (not yet identified) were observed in the ICP-MS trace when plants were exposed to 0.67 µmol As l–1 or more, whereby the number of species increased with concentration (Table 1). No PC complexes of DMA(V) were detected. Generally, the concentrations of all complexes increased with increasing As concentration, but at 66.7 µmol As l–1 the concentrations of As(III)–(PC2)2, As(III)–PC3, and MA(III)–PC2 were lower than at 13.3 µmol As l–1. The concentration of GS–As(III)–PC2, the sum of the unknown complexes and the unbound arsenic in root levelled off. Up to an exposure concentration of 13.3 µmol As l–1 all other identified complexes behaved similarly.
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Considering the whole plant, GS–As(III)–PC2 did increase linearly and the other complexes followed an exponential curve to maximum. The ratio of bound to unbound As followed an equation describing a combination of an exponential increase followed by an exponential decrease starting at concentrations below 6.6 µmol As l–1 (Fig. 4).
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Influence of As(V) concentration on GSH and PC production
Oxidized PC2 was the dominant form of this peptide in all plant parts when not complexed by As. Significant amounts of reduced PC2 were only detected in roots at all concentrations and in leaves of plants exposed to less than 1.3 µmol As l–1, stems contained nearly no reduced PC2 at all. In roots, reduced PC2 increased linearly with exposure, whereas the amount of oxidized PC2 increased exponentially to a maximum. The signal for oxidized PC2 in leaves increased linearly with increasing As concentration up to 13.3 µmol As l–1. The ratio of reduced to oxidized PC2 steadily increased in roots (0.005 to 0.4 at 66.7 µmol As l–1), while in leaves it fell from 0.3 to 0.001.
Reduced glutathione in roots initially increased steeply (about 3.5-fold at 0.13 µmol As l–1 compared with control plants), but with higher As concentrations the increase relative to control plants was only about 2-fold and stayed constant. The amount of oxidized GSH in roots declined steeply up to 1.3 µmol As l–1 and stayed constant thereafter. The ratio of reduced to oxidized GSH in roots and leaves increased with As concentration.
De novo methylation of As(V) by H. annuus
Methylated arsenic species (DMA(V) and MA(V)) were measured after one-time exposure to As(V). DMA(V) is the major methylated arsenic species in H. annuus. MA(V) was detected in some samples, but its overall concentration was very low. Any methylation in roots occurred rapidly and then ceased (Fig. 2). In shoots, the behaviour was similar during the first 8 d. After 16 d the methylation seemed to increase again. Overall, the amount of methylated arsenic was small (less than 1%). The amount of DMA(V) in shoots followed an exponential curve, whereas in roots no correlation with time was detectable (similar for MA(V)).
As toxicity
Root growth (length and weight) of H. annuus was not greatly influenced (95±5% of control) by exposure to low concentrations of As(V) (0–10 µmol As l–1). Growth reduction of 50% and more was observed at As(V) concentrations of more than 15 µmol As l–1 (Fig. 5). Using a logistic four parameter equation the EC50 concentration was 21.6±2.8 µmol l–1, with maximum effect seen at 103±4.6 µmol l–1.
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Discussion |
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In this paper, the concentration-dependent uptake and As–PC complex formation by H. annuus were studied. The nature of the As–PC complexes formed in roots was not strongly affected by exposure time and exposure concentration. Also, As was not translocated between different plant parts of H. annuus once exposure ceased, a similar effect was observed by Sneller et al. (1999) for Silene vulgaris. More than half of the total arsenic taken up and, similarly, more than half of the As–PC complexes at concentrations below 13.3 µmol As l–1, were found in the roots. Reina et al. (2005) found that hydroponically grown Lupinus albus L., exposed to similar As concentrations, retained more than 95% of the arsenic in the roots, but the course of uptake into roots (calculated as µmol As g–1 plant) versus exposure was similar in H. annuus and L. albus. Analogous observations were made for other plant species like Cytisus striatus (Bleeker et al., 2003), Silene vulgaris (Sneller et al., 1999), and Holcus lanatus (Hartley-Whitaker et al., 2002). These plants seem to retain most of the As in the roots, which is in contrast to As hyperaccumulators which export larger amounts toward the shoot (Meharg and Hartley-Whitaker, 2002). Nevertheless, exposure to As concentrations above the EC50 value for a particular species increased the relative amount of As found in the shoot system, for example, in our study H. annuus exposed to 66.7 µmol As l–1 retained 50% of the As in the roots in contrast to 85% when exposed to 0.67 µmol As l–1. Reina et al. (2005) observed a similar effect in Lupinus albus L. At the same time the As uptake relative to available As was reduced at higher external As concentrations. This reduction did run parallel with the relative toxicity index (RTI), but started at lower As concentrations (<1.3 µmol As l–1). Sneller et al. (1999) observed a similar effect in Silene vulgaris.
According to the RTI (determined from root growth of 7-d-old seedlings) the EC50 value for H. annuus was in the range of 22 µmol As l–1, but reduced As uptake rate at concentrations over 1.3 µmol As l–1 could indicate that As started to have toxic effects on H. annuus at concentrations 20 times lower than determined by root growth. Another explanation might be that uptake regulation starts to have an effect. Analogous conclusions could be drawn from the ratio of PC-bound As versus unbound As in the roots, which reached its highest level at 1.3 µmol As l–1. The reduction of this ratio at higher As concentrations could show that As is interfering negatively with peptide and/or protein synthesis.
Considering the amount of individual As–PC complexes the influence of As toxicity on PC synthesis was more similar to the RTI results. Most of them (As(III)–(PC2)2, MA(III)–PC2 and As(III)–PC3) followed an exponential increase reaching maximum level at 13.3 µmol As l–1. Sneller et al. (1999) observed a similar trend in Silene vulgaris for total PCs. Only the GS–As(III)–PC2 complex showed linear behaviour with increasing exposure over the whole concentration range in H. annuus.
Sneller et al. (1999, 2000) and Reina et al. (2005) described a positive relationship between PC-concentrations as determined by Ellmann's reagent or monobromobimane and As exposure in roots. Both authors describe that, at high As concentrations, the amount of PCs present in the root is reduced, similar to our finding that the main As–PC complexes were reduced at 66.7 µmol As l–1. The rapid increase of PC2 at low exposure concentrations showed the strong relationship between PC-synthesis and As exposure as described by Schat et al. (2002) for other plant species. Our ES-MS data equally showed that not all synthesized PC2 was used for either binding to As or for chain-elongation to PC3. PC2 has, at least in H. annuus, a role similar to glutathione, namely that it can serve as reducing agent and therefore potentially protect other more essential systems from damage due to As. This cannot be observed using traditional UV or fluorescence-methods for quantification of PCs as used by Sneller et al. (1999) since PC2 prefers the intramolecular oxidation, so that no reaction with either Ellmann's reagent or monobromobimane can occur. Glutathione synthesis on the other hand did not seem to increase drastically in H. annuus which was equivalent to results found in Silene vulgaris by Sneller et al. (1999).
Methylation of inorganic arsenic did not appear to play a major role in the detoxification of inorganic arsenic by H. annuus. Methylation after one month was less than 1%, similar to results found by Kuehnelt et al. (2000) for other terrestrial plants. Additional evidence that methylation is unimportant in higher terrestrial plants comes from the findings of Nissen and Benson (1982), who specifically studied the factors influencing methylation of arsenic by plants and found that unless the nitrogen supply is strongly limited (which it was not in this experimental set-up) methylation in terrestrial plants does not occur. That the small amount of methylated arsenic was, in fact, methylated by bacteria in the surrounding soil cannot be excluded from our data, but is unlikely since it would require that As is re-excreted into the soil, methylated by bacteria, and then taken up again.
Generally, it can be said that parameters of the cellular metabolism, like PC-synthesis, showed the negative influence of arsenic at much lower concentrations than could be detected by comparing general plant growth parameters, like root growth or weight.
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Acknowledgements |
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The authors thank BBSRC for financial support while KF thanks the Erasmus programme.
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Abbreviations |
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ICP-MS, inductively coupled plasma mass spectrometer; ES-MS, electrospray mass spectrometer; HPLC, high pressure liquid chromatography; PC, phytochelatin; As(III), arsenite; As(V), arsenate; As–PC, arsenic–phytochelatin complex; FW, fresh weight; DMA(V), dimethylarsinic acid; MA(V), monomethylarsonic acid; RTI, root tolerance index; GSH, reduced glutathione; GSSG, oxidized glutathione; PC2, phytochelatin-2 [(
-Glu-Cys)2-Gly]; PC3, phytochelatin-3 [(-Glu-Cys)3-Gly]; As(III)–(PC2)2 (1) and As(III)–(PC2)2 (2), isomers of complex of As(III) with two (-Glu-Cys)2-Gly molecules; GS–As(III)–PC2, complex of As(III) with GSH and (-Glu-Cys)2-Gly; As(III)–PC3, complex of As(III) with (-Glu-Cys)3-Gly; MA(III)–PC2, complex of MA(III) with (-Glu-Cys)2-Gly; MA(III), methylarsonous acid. |
References |
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