Ecotoxicology (2009) 18:110–121 DOI 10.1007/s10646-008-0264-3
Bioaccumulation and physiological effects of mercury in Pteris vittata and Nephrolepis exaltata Jian Chen Æ Safwan Shiyab Æ Fengxiang X. Han Æ David L. Monts Æ Charles A. Waggoner Æ Zhimin Yang Æ Yi Su
Accepted: 22 August 2008 / Published online: 3 September 2008 Ó Springer Science+Business Media, LLC 2008
Abstract Anatomical, histochemical and biochemical approaches were used to study mercury uptake and phytotoxicity as well as anti-oxidative responses in two species of ferns [Chinese brake fern (Pteris vittata) and Boston fern (Nephrolepis exaltata)], grown in a hydroponic system. The roots of both cultivars accumulated large amounts of mercury, but exhibited limited mercury translocation to shoots. Mercury exposure led to more pronounced phytotoxicity accompanied by stronger oxidative stress in the shoots of P. vittata than in N. exaltata. N. exaltata established a more effective anti-oxidative system against mercury-induced oxidative stress than did P. vittata. The activity of anti-oxidative enzymes (superoxide dismutase, catalase and glutathione reductase) increased. The reduced ascorbate (ASA) and oxidized ascorbate (DHA) are regulated. Mercury exposure led to an increase in the concentration of glutathione (GSH) in both fern species. The present study suggests that N. exaltata is more tolerant to mercury exposure than P. vittata, which has been also J. Chen Z. Yang Department of Biochemistry & Molecular Biology, College of Life Science, Nanjing Agricultural University, Nanjing 210095, People’s Republic of China J. Chen S. Shiyab F. X. Han (&) D. L. Monts C. A. Waggoner Y. Su Institute for Clean Energy Technology (ICET), Mississippi State University, Starkville, MS 39759, USA e-mail:
[email protected] F. X. Han Department of Plant and Soil Sciences, Mississippi State University Mississippi State, Starkville, MS 39762, USA D. L. Monts Y. Su Department of Physics & Astronomy, Mississippi State University, Starkville, MS 39762, USA
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reported to be more tolerant to arsenic exposure. N. exaltata may thus have potential for phytostabilization of soils or phytofiltration of waste water contaminated with mercury. Keywords Mercury Phytotoxicity Oxidative stress Pteris vittata Nephrolepis exaltata Phytoremediation
Introduction Mercury (Hg) is one of the major toxic metals because it bioaccumulates and biomagnifies in animal and human bodies by way of the food chain (Suszcynsky and Shann 1995). Millions of tons of mercury have been released and accumulated in the global environment since the beginning of the Industrial Revolution (Han et al. 2002). The main sources of mercury pollution include mining of gold and silver, the coal industry, untreated discarded batteries, and industrial waste disposal (Pilon-Smith and Pilon 2000). Mercury can also enter into agricultural soil by anthropogenic activities, such as fertilizers, sludge, pesticides, lime, and manure (Han et al. 2002). Mercury in soil exists in many forms, including elemental mercury (Hg0), ionic mercury (Hg2?), methyl mercury (MeHg), mercury hydroxide (Hg(OH)2), and mercury sulfide (HgS). Hg2? is the predominant toxic form of mercury (Heaton et al. 2005). Mercury may be directly taken up by plants and may subsequently lead to toxic reactions (Han et al. 2006). Phytoremediation is a cost-effective technology that utilizes plants to remove, transform, or stabilize contaminants, including organic pollutants and toxic metals in water, sediments, or soils (Cherian and Oliveira 2005). Phytoremediation has been accepted and utilized widely
Bioaccumulation and physiological effects of mercury
because it is cost-effective; permanently removes the pollutant, and protects nature. Over 400 plant species have been identified as natural metal hyperaccumulators (Brooks 1998; Reeves and Baker 2000). For example, Alyssum murale, Thlaspi rotundifolium, Pteris vittata and Sesbania drummondii are known for their ability to hyperaccumulate Ni, Zn, As, and Pb, respectively (Ma et al. 2001; Rasico 1977; Sahi et al. 2002; Severne and Brooks 1972). However, no natural hyperaccumulator for mercury has been indentified to date. Although the detailed mechanism of oxidative stress in plants caused by heavy metal exposure is not yet fully understood, it is recognized that the establishment of an anti-oxidative system is an important mechanism for plants to respond to heavy metal stress. Mercury triggers oxidative stress by inducing the production of hydrogen peroxide (H2O2), lipid peroxides, and reactive oxygen species (ROS) in many plants, such as tomato, cucumber, and alfalfa (Cho and Park 2000; Cargnelutti et al. 2006; Zhou et al. 2007, 2008). Plant cells have antioxidants (like a-tocopherol, glutathione, and ascorbate) and anti-oxidative enzymes (such as superoxide dismutase (SOD), guaiacol peroxidase (POD), catalase (CAT) and glutathione reductase (GR), which participate in scavenging ROS (Halliwell 1982). Plants can adapt to oxidative stress through the production of these antioxidants and anti-oxidative enzymes. For example, mercury stress induces a significant increase in the activity of anti-oxidative enzymes in Sesbania drummondii and Zea mays (Israr et al. 2006; ´ lvarez et al. 2006). Compared with arsenic-sensitive Rella´n-A fern species, arsenic-hyperaccumulative fern species, such as P. vittata, have higher concentration of antioxidants and greater activity of anti-oxidative enzymes (Cao et al. 2004). In addition, the detoxification of heavy metals in plants may involve chelating toxic metals by phytochelatins (PCs). PCs are heavy-metal-binding peptides with a repeat unit of (c-glu-cys)n-Gly that are synthesized from glutathione (GSH) (Zenk 1996). The relationship between glutathione metabolism and phytochelatin synthesis has been shown clearly by kinetic experiments in vivo (Scheller et al. 1987). A previous study has shown that mercury stress leads to an increase of glutathione and nonprotein thiols levels (Zhou et al. 2007), which suggests a possible linkage between glutathione and the defense system against heavy metal stress. Because it has been identified an arsenic hyperaccumulator, a considerable number of studies have been conducted to investigate the detoxification mechanism of Chinese brake fern (Pteris vittata) after arsenic stress (Cao et al. 2004; Ma et al. 2001; Singh et al. 2006; Zhao et al. 2003). Since no natural hyperaccumulator plant has yet
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been found for mercury, we investigated the oxidative responses of two fern cultivars [Chinese brake fern (P. vittata) and Boston fern (N. exaltata)] under mercury stress in a hydroponic system. The objectives of this study were to investigate (1) the uptake of mercury by two species of fern; (2) the phytotoxicity of mercury to the two ferns; and (3) the oxidative stress and anti-oxidative defense responses of the two ferns to mercury. Ultimately, the present study aimed to examine the potential use of the two fern species in phytoremediation of mercury contaminated soils and water. In addition, since mercury toxicity to plant roots has been well documented in the literature, our primary focus is on the response of plant shoots to mercury stress.
Materials and methods Plant growth and experiment design Chinese brake fern (P. vittata), 3 months old, were obtained from Edenspace Systems (Dulles, VA, USA). Boston fern (N. exaltata)] 3 months old, were obtained from Walmart (Starkville, MS, USA). The roots of two ferns were washed with distilled water and then transferred into a modified full-strength Hoagland solution containing 33.95 mg l-1 KH2PO4, 19.93 mg l-1 NH4NO3, 126.44 mg l-1 KNO3, 60.19 mg l-1 MgSO4, 164.1 mg l-1 Ca(NO3)2, 0.71 mg l-1 H3BO3, 0.02 mg l-1 CuSO4, 1.33 mg l-1 FeSO4 7H2O, 0.604 mg l-1 MnSO4 H2O, 0.004 mg l-1 MoO3, and 0.055 mg l-1 ZnSO4. The pH of the nutrition was 5.5, which has been shown to be suitable for fern growth (Srivastava et al. 2005). Mercury treatment started after 3 days by adding Hg(NO3)2 H2O to the solution. The mercury treatments included the control, 5, and 20 mg Hg l-1. The free mercury concentrations in hydroponic solutions were validated as 0, 4.11, and 16.7 mg Hg l-1 (Table 1). All the treatment groups along with the control were arranged in a completely randomized design. Each group consisted of three replicates. Three and seven days after the mercury treatment, fresh leaf samples were collected and weighed for microscopy study and biochemical measurements. At the end of the treatment, leaves and roots were harvested, and the fresh weights (FW) of shoots were determined. The samples were dried in a forced draft oven for 48 h at 80°C for determination of dry weights (DW). The relative water content (RWC) was calculated using RWC (%) = [(FW - DW)/FW] * 100. Plant roots were washed with 1% (v/v) HCl for several seconds, followed by washing three times with distilled water in order to remove the mercury adhering to the surface of the roots.
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Table 1 Concentrations of mercury (in mg kg-1 DW, DW = dry weight) in shoots and roots of P. vittata and N. exaltata Hg treatment (mg l-1)
Measured Hg in solution (mg l-1)
Hg concentration in shoot (mg kg-1 dw)
Hg concentration in shoot (mg kg-1 dw)
Ratios of Hg concentration in root
P. vittata
P. vittata
P. vittata
N. exaltata
1.9 ± 0.5a
N. exaltata
N. exaltata
0
0
0.7 ± 0.0a
0.5 ± 0.1a
1.2 ± 0.2a
2.7 ± 0.6a
2.5 ± 0.8a
5
4.11
4.5 ± 1.6bB
1.0 ± 0.2bA
1298 ± bA
1117 ± 210bA
303 ± 95bA
1240 ± 411cB
20
16.7
5.8 ± 1.6bA
3.1 ± 0.5bA
3534 ± cA
2645 ± 437cA
616 ± 46cA
850 ± 5.8bB
-1
Plants were exposed to 0, 4.11, or 16.7 mg l Hg for 7 days. Data are means with standard errors (n = 3). Means followed by the same letter were not significantly different at P \ 0.05 (a–c denote significance within a species among different mercury treatments; A–B denote significance between the two species)
Mercury analysis in plant tissue
Determination of enzyme activities
About 0.15 g of dried plant sample was mixed with 1.5 ml concentrated HNO3 in a test tube and covered with Parafilm for 24 h. The samples were digested with H2O2 (30% w/v). The digested solution was analyzed for mercury concentration using cold vapor atomic absorption spectrometry (CVAAS, FIMS-100) and inductively coupled plasma-atomic emission spectrometry (ICP-AES, Optima 4300 DV). And calibration was performed by using standard Hg solution (Han et al. 2006).
About 0.05 g of leaf tissue was homogenized in 1.5 ml of ice-cooled phosphate buffer (50 mM, pH 7.0, containing 1 mM ethylenediamine tetra-acetic acid (EDTA) and 1% w/v insoluble polyvinylpyrrolidone). The homogenate was centrifuged at 15,000 g for 10 min at 4°C. The supernatant was used as the crude extract for the assay of enzyme activities. Activity of superoxide dismutase (SOD; EC 1.1.5.1.1) was assayed by measuring the inhibition of photochemical reduction of nitro-blue tetrazolium (NBT) (Beauchamp and Fridovich 1971). The reaction mixture (3 ml) contained 50 mM potassium phosphate buffer (pH 7.8), 13 mM methionine, 2 lM riboflavin, 75 lM NBT, 0.1 mM EDTA and 30 ll enzyme extract. The absorbance of the solution was measured at 560 nm. One unit of SOD was defined as the amount of enzyme that causes half-maximal inhibition of the NBT reduction under assay conditions. Activity of catalase (CAT; EC 1.11.1.6) was assayed by measuring the decomposition of hydrogen peroxide. About 50 ll enzyme extract was added to the reaction mixture containing 1 ml phosphate buffer solution (50 mM, pH 7.0) and 0.1% H2O2. The decrease of the absorbance at 240 nm was recorded. Activity was calculated using an extinction coefficient of 0.036 mM-1 cm-1. One unit of CAT activity was defined as the amount required to decompose 1 lmol of hydrogen peroxide min-1 mg-1 protein under assay conditions (Beers and Sizer 1952). Glutathione reductase (GR; EC 1.8.5.1) activity was determined by transformation of oxidized glutathione (GSSG) to glutathione (GSH) (Sgherri et al. 1994). One ml assay mixture contained 200 mM potassium phosphate buffer (pH 7.5), 0.2 mM Na2EDTA, 1.5 mM MgCl2, 0.5 mM GSSG, 50 lM triphosphopyridine nucleotide (NADPH), and 200 ll enzyme extract. The reaction was initiated by addition of NADPH and the decrease in absorbance at 340 nm was recorded. Corrections were made for non-enzymatic oxidation of NADPH by recording the decrease at 340 nm without adding GSSG to the assay mixture. The enzyme activity was calculated from the
Determination of lipid peroxidation The concentration of thiobarbituric acid reactive substances (TBARS) was determined as an indicator of the level of lipid peroxidation in the leaves. About 0.05 g of fresh leaf sample was ground with 1.5 ml of 0.1% (w/v) trichloroacetic acid (TCA). The homogenate was centrifuged at 10,000 g for 10 min and 0.5 ml of the supernatant was mixed with 2 ml of 20% (w/v) TCA-containing 0.5% (w/v) 1,3-diethyl-2-thiobarbituric acid (TBA). The mixture was heated at 90°C for 20 min and then cooled in ice. The mixture was then centrifuged at 10,000 g for 5 min. The absorbance of the supernatant was measured at 532 nm and at 600 nm. The TBARS content was calculated using an extinction coefficient of 155 mM-1 cm-1 (Ohkawa et al. 1979). Determination of H2O2 Approximate 0.1 g of leaf sample was homogenized with a mortar and pestle in 1.5 ml phosphate buffer (50 mM, pH 6.5), containing 1 mM hydroxylamine. After centrifugation at 10,000 g for 10 min, 0.5 ml supernatant was mixed with 1.5 ml 20% (v/v) H2SO4, containing 0.1% (v/v) TiCl4. After thorough mixing, the mixture was centrifuged at 10,000 g for 10 min. Then the absorbance of the supernatant was measured at 410 nm. The content of H2O2 was calculated using an extinction coefficient of 0.28 lM-1 cm-1 (Jana and Choudhuri 1981).
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initial rate of the reaction after subtracting non-enzymatic oxidation by using an extinction coefficient of 6.2 nM-1 cm-1. One unit of GR activity is defined as the amount required to decompose 1 lmol of NADPH min-1 mg-1 protein under assay conditions.
curve was developed based on GSH in the range 0–50 lM to convert the measured absorbance to concentration (Smith 1985).
Determination of non-enzymatic antioxidants
Protein in samples was quantified by the Bradford method (Bradford 1976), using bovine serum albumin as standard.
Leaf samples were prepared for ascorbate (ASA) and glutathion (GSH) analyses by homogenizing 1 g of leaf material (fresh weight) in 10 ml of cold 5% (w/v) metaphosphoric acid (Gossett et al. 1994). The homogenate was centrifuged at 22,000 g for 15 min at 4°C, and the supernatant was collected for analyses of ascorbate and glutathione. Reduced ascorbate (ASA) and oxidized ascorbate (DHA) were measured according to Law et al. (1983). Total ascorbate was determined after reduction of DHA to ASA with 1,4-dithiothreitol (DTT). The concentration of DHA was estimated from the difference between total ascorbate and ASA. The reaction mixture for total ascorbate contained 0.3 ml aliquot supernatant, 0.75 ml 150 mM phosphate buffer (pH 7.4) containing 5 mM EDTA, and 0.5 ml 10 mM DTT. After incubation for 10 min at room temperature, 0.15 ml 0.5% N-ethylmaleimide was added to remove excess DTT. ASA was determined in a similar reaction mixture, except that 0.3 ml H2O was added instead of DTT and N-ethylmaleimide. Color developed in both reaction mixtures after addition of the following reagents: 0.6 ml of 10% (w/v) TCA, 0.6 ml of 44% (v/v) ortho-phosphoric acid, 0.6 ml 4% (w/v) a,a0 dipyridyl in 70% (v/v) ethanol, and 0.3% (w/v) FeCl3. After vortex mixing, the mixture was incubated at 40°C for 40 min and the absorbance at 525 nm was recorded by using Genesys 6 spectrophotometer (Thermo Electron Corporation, Waltham, MA, USA). A standard curve of ASA in the range 0–100 lg ml-1 was prepared to convert the measured absorbance to concentration. Redox status was calculated by the ratio of the amount of ASA to the total amount of ASA plus DHA. For GSH assay, one ml aliquot of the supernatant was neutralized with 1.5 ml of 0.5 M phosphate buffer (pH 7.5) after which 50 ll H2O was added. This sample was used for assay of total glutathione. Glutathione content was measured in a 3 ml reaction mixture containing 0.2 mM NADPH, 100 mM phosphate buffer (pH 7.5), 5 mM EDTA, 0.6 mM 5,50 -dithiobis(2-nitrobenzoic acid) (DTNB), and 3 units of GR. The reaction was started by adding 0.l ml of the extract sample obtained as described above. The reaction rate was monitored by measuring the change in absorbance at 412 nm for 1 min. The absorbance was record by using Genesys 6 spectrophotometer (Thermo Electron Corporation, Waltham, MA, USA). A standard
Determination of protein content
Scanning electron microscopy (SEM) The plant samples were cut into small pieces and immediately fixed in 2.5% (v/v) glutaraldehyde in 0.05 M potassium phosphate buffer (pH 7.1) for 8 h. The samples were dehydrated in an ethanol series. The samples were sealed in Parafilm, frozen in liquid nitrogen, and fractured transversely using a pre-cooled knife. The cryofractured specimens were critical-point dried using carbon dioxide, mounted on stubs, and coated with gold–palladium. All materials were observed with a LEO SEM (Su et al. 2005). Transmission electron microscopy (TEM) The samples were cut into small pieces and fixed in 2.5% glutaraldehyde in 0.05 M potassium phosphate buffer (pH 7.1) for 8 h, and post-fixed with OsO4. The samples were dehydrated in an ethanol series (95%, v/v) and embedded in Spurrs epoxy resin. Ultrathin sections were obtained using an ultramicrotome and stained with uranyl acetate and basic lead citrate for observation using a JEOL TEM (Su et al. 2005). The average diameter and the number of vascular cells were used for quantifying cell structure changes in TEM micrographs under mercury stress. Histochemical detection of H2O2 Histochemical detection of H2O2 was performed in the fern leaves using 3,3-diaminobenzidine (DAB) as a substrate. The treated fern plants were excised at the base of the stems with a razor blade and supplied through the cut stems with 1 mg ml-1 solution of DAB (pH 3.8) for 6 h. The leaves were immersed in boiling ethanol (v/v 95%) for 20 min to remove the green background. This allowed the deep brown polymerization product (reaction of DAB and H2O2) to be clearly visualized and photographed (OrozcoCa´denas and Ryan 1999). Statistical analysis Each result shown in figures is the mean of three replicated treatments. The significance of differences among treatments and between the two fern species was statistically
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evaluated by standard deviation and Student’s t-test methods, using Microsoft Excel 2000 (Microsoft, USA).
Results Uptake of mercury P. vittata and N. exaltata were exposed to 0, 4.11, and 16.7 mg l-1 mercury for up to 7 days. Mercury concentrations increased in both shoots and roots of these two fern species with an increase in mercury concentration in the growth medium. More mercury was accumulated in the roots than in the shoots of both fern species, especially at 16.7 mg l-1 mercury exposure level (Table 1). N. exaltata had much higher ratios (850–1240) of mercury concentrations in roots to shoots than did P. vittata (303–616) at the two mercury exposure levels (Table 1). There were no significant differences in mercury concentration in the roots between the two fern species. Compared to N. exaltata, P. vittata accumulated more mercury in shoots. Physiological effects of mercury in ferns Compared to the control, the relative water content (RWC) in the shoots of P. vittata decreased by 21% and 29% at 4.11 and 16.7 mg l-1 mercury levels, respectively (Table 2). However, RWC in the shoots of N. exaltata did not show a significant change at the mercury exposure levels investigated (Table 2). Severe visual toxic symptoms (such as withering, chlorosis and falling of leaves) appeared in P. vittata, especially at 16.7 mg l-1 mercury level (Fig. 1a); in contrast, no toxic symptoms were observed for N. exaltata (Fig. 1b). Mercury exposure led to leaf burn in P. vittata (Fig. 1c). The electron microscopy study also indicated different toxic effects of mercury on the cellular structure in leaves of P. vittata and N. exaltata. The SEM micrographs showed changes of the vascular cells of the leaf samples (Fig. 2). Exposure to 16.7 mg l-1 mercury for 7 days resulted in a loss of cell shape, decreases in the intercellular spaces, and shrinkage of vascular bundle in P. vittata (Fig. 2b) compared to the control (Fig. 2a). Some
precipitates filled in the intercellular spaces in leaves of P. vittata after mercury treatment (Fig. 2b). For N. exaltata, changes in vascular cell structure were not significant for mercury treatment (Fig. 2d) compared to the control (Fig. 2c). Furthermore, after plant exposure to 16.7 mg l-1 mercury for 7 days, the TEM micrographs showed a significant decrease in the amount of chloroplast in leaf cells of P. vittata (Fig. 3b) compared to the control (Fig. 3a). For P. vittata, 16.7 mg l-1 mercury treatment (Fig. 3d) led to a significant breakdown of thylakoid inside the chloroplasts of the leaf cells compared to the control (Fig. 3c). We did not find significant damage of the chloroplast in N. exaltata under mercury exposure. Effect of mercury on hydrogen peroxide accumulation Exposure to mercury induced a significant accumulation of hydrogen peroxide (H2O2) in the shoots of P. vittata. After exposure of mercury for 3 days, H2O2 concentrations in the shoots of P. vittata increased by 87% and 126% compared to control at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 4a). After exposure of mercury for 7 days, H2O2 concentrations in the shoots of P. vittata increased by 229% and 290% compared to control at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 4a). However, H2O2 concentration in N. exaltata did not show a significant change at either mercury level (Fig. 4b). Histochemical staining also indicated H2O2 accumulation in vivo. H2O2 can react with 3,3-diaminobenzidine (DAB) to appear pink, which can be used as an indicator of H2O2 production in vivo. Compared to the control, more pink color appeared inside of nervation in leaves of P. vittata after exposure to 16.7 mg l-1 mercury for 7 days (Fig. 4c). Effect of mercury on lipid peroxidation Lipid peroxidation reflected the toxic effects of mercury on the cell membrane in tissues. The concentration of TBARS in the shoots of P. vittata increased by 390% at 16.7 mg l-1 mercury level for 3 days (Fig. 5a). After 7 days of exposure to mercury, P. vittata showed a further increase by 230% and 520% times in the concentration of TBARS at 4.11 and 16.7 mg l-1 mercury levels,
Table 2 Effect of mercury on the dry weight (DW) and relative water content (RWC) in shoots of P. vittata and N. exaltata Hg treatment (mg l-1) (measured)
DW (g) P. vittata
RWC (%) N. exaltata
P. vittata
N. exaltata
0
1.5 ± 0.3a
1.2 ± 0.2a
74.1 ± 2.7b
84.5 ± 0.1a
4.11
1.6 ± 0.2a
1.3 ± 0.3a
58.5 ± 1.4a
84.3 ± 0.6a
1.4 ± 0.2a
1.1 ± 0.4a
52.5 ± 6.5a
83.5 ± 0.9a
16.7 -1
Plants were exposed to 0, 4.11, or 16.7 mg l Hg for 7 days. Data are means with standard errors (n = 3). Means followed by the same letter were not significantly different at P \ 0.05 (a–c denote significance within a species among different mercury treatments)
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Fig. 1 Effect of mercury on shoots of P. vittata (a) and N. exaltata (b). Plants were exposed to 0, 4.11, or 16.7 mg l-1 Hg for 7 days. (c) Leaf burns in P. vittata under 16.7 mg l-1 Hg exposure. Arrows show mercury-induced toxic effects
Fig. 2 SEM micrographs of fern leaf cellular structure. Plants were exposed to 16.7 mg l-1 mercury for 7 days: the control (a) and 16.7 mg l-1 Hg treatment (b) of P. vittata; the control (c) and 16.7 mg l-1 Hg treatment (d) of N. exaltata. Arrows show the deformation of cell shape and the shrinkage of vascular bundle in P. vittata induced by mercury
respectively (Fig. 5a). Mercury exposure did not significant change the concentration of TBARS in the shoots of N. exaltata. However, TBARS level in shoots of N. exaltata did slightly increase with the time of mercury treatment (Fig. 5b).
Effect of mercury on antioxidant enzymes Compared to the control, there was no significant change in SOD activity in the shoots of P. vittata after mercury exposure (Fig. 6a). For N. exaltata, SOD activity showed a
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116 Fig. 3 TEM micrographs of chloroplast (as indicated with arrows) in the leaf cells of P. vittata from the control (a) and 16.7 mg l-1 mercury treatment for 7 days (b). The micrographs were zoomed in to show the effects of mercury on thylakoid (as indicated with arrows) inside the chloroplast at the control (c) and 16.7 mg l-1 Hg treatment for 7 days (d)
Fig. 4 Effect of mercury on the concentration of hydrogen peroxide (H2O2) (in nmol g-1 FW, FW = fresh weight) in the shoots of P. vittata (a) and N. exaltata (b). Plants were exposed to 0, 4.11, or 16.7 mg l-1 Hg for 3 and 7 days, respectively. Error bars represent standard deviation of three separate experiments. Means followed by the same letter were not significantly different at P \ 0.05 (a–c denote significance within a species among different mercury treatments). (c) Histochemical stain of H2O2 in vivo in leaves of P. vittata. Plants were exposed to 0 and 16.7 mg l-1 Hg for 7 days. Plants were harvested and stained with 3,3diaminobenzidine (DAB) solution (For detailed description, see ‘‘Materials and Methods’’ section) and immediately photographed
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Fig. 5 Effect of mercury on the concentration of TBARS (in nmol g-1 FW, FW = fresh weight) in the shoots of P. vittata (a) and N. exaltata (b). Plants were exposed to 0, 4.11, or 16.7 mg l-1 Hg for 3 and 7 days, respectively. Error bars represent standard
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deviation of three separate experiments. Means followed by the same letter were not significantly different at P \ 0.05 (a and b denote significance within a species among different mercury treatments)
Fig. 6 Effect of mercury on the activity of superoxide dismutase (SOD) (in unit mg-1 protein), catalase (CAT) (in lmol min-1 mg-1 protein), and glutathione reductase (GR) (in lmol min-1 mg-1 protein) in shoots of P. vittata (a, c, and e) and N. exaltata (b, d, and f). Plants were exposed to 0, 4.11, or 16.7 mg l-1 Hg for 3 and 7 days, respectively. Error bars represent standard deviation of three separate experiments. Means followed by the same letter were not significantly different at P \ 0.05 (a and b denote significance within a species among different mercury treatments)
significant increase by 85% and 76% at 4.11 and 16.7 mg l-1 mercury levels for 7 days, respectively (Fig. 6b). Compared to the control groups, mercury exposure did cause a decrease in the activity of CAT in P. vitatta (Fig. 6c). In contrast, CAT activities in the shoots of N. exaltata increased by 27% and 67% at 16.7 mg l-1 mercury levels after 3 and 7 days, respectively (Fig. 6d). Mercury treatment had no significant effect on
the activity of GR in P. vittata, except for an increase in GR activity at 4.11 mg l-1 mercury treatment for 7 days (Fig. 6e). GR activity in N. exaltata increased by 120% at 16.7 mg l-1 mercury level after 3 days of mercury treatment (Fig. 6f). After 7 days of exposure to mercury, GR activity in N. exaltata showed a significant increase by 130% and 108% at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 6f).
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However, compared to the control, N. exaltata did not show significant change in DHA concentration at either mercury level (Fig. 8b). Compared to the control, the ratios of ASA/ (DHA ? ASA) in the shoots of P. vittata decreased by 7% and 14% at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 8a). In contrast, N. exaltata showed an increase by 11 - 13% in the ratios of ASA/(DHA ? ASA) at 4.11 and 16.7 mg l-1 mercury levels (Fig. 8b).
Discussion Fig. 7 Effect of mercury on the concentration of glutathione (GSH) (in lmol g-1 FW, FW = fresh weight) in the shoots of P. vittata and N. exaltata. Plants were exposed to 0, 4.11, or 16.7 mg l-1 Hg for 7 days. Error bars represent standard deviation of three separate experiments. Means followed by the same letter were not significantly different at P \ 0.05 (a and b denote significance within a species among different mercury treatments)
Effect of mercury on non-enzymatic antioxidants After 7 days of mercury exposure, the concentration of GSH in the shoots of P. vittata increased by 356% and 249% at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 7). N. exaltata showed an increase by 67% and 48% in GSH concentration at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 7). After 7 days of mercury treatment, the reduced ascorbate (ASA) concentration in shoots of P. vittata increased by 25% at 4.11 mg l-1 mercury level, but decreased by 24% at 16.7 mg l-1 mercury level as compared to the control (Fig. 8a). Shoots of N. exaltata showed an increase by 44% and 22% in ASA concentration at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 8b). For the oxidized ascorbate (DHA), P. vittata increased by 20-fold and 13-fold compared to the control at 4.11 and 16.7 mg l-1 mercury levels, respectively (Fig. 8a).
Fig. 8 Effect of mercury on the concentration of oxidized ascorbate (ASA) (in lmol g-1 FW, FW = fresh weight), reduced ascorbate (DHA) (in lmol g-1 FW, FW = fresh weight) and the ratio of ASA/ (ASA ? DHA) (indicated with the number on the top of columns) in the shoots of P. vittata (a) and N. exaltata (b). Plants were exposed to
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The present study focused on mercury uptake and cellular structural changes of two cultivars of fern, P. vittata and N. exaltata, as well as anti-oxidative responses caused by mercury toxicity. Chinese brake fern, P. vittata, has been found to be a natural hyperaccumulator plant for arsenic (Ma et al. 2001). Therefore, we also briefly compare the biochemical responses of both ferns to mercury (present study) and their reactions to arsenic (as reported in the literature). The results presented here indicate that large amounts of mercury accumulated in the roots of both fern species (Table 1). Under similar treatments, previous studies reported much lower mercury accumulations in the roots of Bacopa monnieri, tomato, and cucumber (Cargnelutti et al. 2006; Cho and Park 2000; Sinha et al. 1996). On the other hand, a high mercury accumulation (about 2500 mg kg-1 DW) has been reported in the roots of Sesbania drummondii (Cory Rattlebush), which was considered for phytoremediation of a mercury-contaminated environment (Israr et al. 2006). In the present study, the transfer of mercury from roots to shoots was limited in both fern species. In contrast, Ma et al. (2001) suggested that P. vittata could take up a large amount of arsenic via their roots and then transfer most of this to their shoots, which
0, 4.11, or 16.7 mg l-1 Hg for 7 days. Error bars represent standard deviation of three separate experiments. Means followed by the same letter were not significantly different at P \ 0.05 (a–c denote significance within a species among different mercury treatments)
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probably indicates the different translocation mechanism between mercury and arsenic in P. vittata. Relative moisture content (RWC%) has been suggested as an indicator of phytotoxicity after heavy metal stress (Zn and Cr) in Indian mustard and Chinese brake fern (Su et al. 2005). These authors found that RWC was correlated to concentrations and phytotoxicity of heavy metals in plants. RWC in P. vittata shoots decreased significantly under mercury exposure. Since leaf RWC indicates the water status in plant, reduced leaf RWC seems to have disturbed the homeostasis of P. vittata due to the mercury stress. However, leaf RWC of N. exaltata was not suppressed by mercury exposure. The biomass (dry weight) was not significantly different in the mercury treatments compared to the controls (Table 2). This may be explained by the fact that ferns generally grow slowly and the present experiments used a short period of time (7 days). Chlorosis was observed in tomato seedlings under mercury stress (Cho and Park 2000). Lenti et al. (2002) reported that mercury inhibited the activity of the NADPH: protochlorophyllide oxidoreductase (POR) that was responsible for the biosynthesis of chlorophyll. In the present study, chlorosis mainly occurred in mature leaves, but not in young leaves, which suggests the possibility of decomposition of chlorophyll in P. vittata under mercury stress. This possibility is in agreement with the TEM micrographs that show breakdown of thylakoid and chloroplast in the leaves of P. vittata after mercury exposure. This ultrastructural change in the shoots may not be specific to mercury stress, but the result of overall metabolic toxicity. The shrinkage of vascular cells in P. vittata observed in SEM micrographs may be caused by the loss of water in cells after mercury exposure, which is in agreement with the observed decrease in RWC of shoots of P. vittata. The current study indicates that P. vittata is more sensitive to mercury stress than N. exaltata, in contrast to the arsenic study where P. vittata is more tolerant than other fern species, including N. exaltata, under arsenic stress (Srivastava et al. 2005). Mercury-induced cellular oxidative damage in plants has been shown to be due to excessive accumulation of reactive oxygen species (ROS), including free radicals and H2O2 (Cargnelutti et al. 2006; Cho and Park 2000; Zhou et al. 2007). As an important signal molecule in plants, H2O2 is indispensable in regulation of plant growth and resistance, but excessively high concentration of H2O2 coupled with ROS can cause lipid peroxidation by attacking membrane lipids (Apel and Hirt 2004). TBARS formed from the decomposition of certain primary and secondary lipid peroxidation products can be used as an indicator of lipid peroxidation in tissues (David 1990). Statistical analysis showed that mercury exposure led to significant accumulation of H2O2 and TBARS in the shoots of
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P. vittata, but not in N. exaltata. This indicates that, in contrast to N. exaltata, mercury induced severe oxidative stress in P. vittata. As an arsenic-hyperaccumulator plant, P. vittata produces less TBARS and H2O2 under arsenic stress compared to arsenic-sensitive fern species (Singh et al. 2006). It seems that P. vittata has different responses to arsenic and mercury stress. High levels of lipid peroxidation and H2O2 accumulation can damage and cause degradation of the chloroplast (Singh et al. 2006). This is in agreement with our observation that degradation of the chloroplast was induced in leaves of P. vittata under mercury stress, as indicated by TEM micrographs. In addition, increased levels of ROS could trigger a programmed cell death pathway (Mitsuhara et al. 1999), which seems to contribute to the death of leaf cells in P. vittata, as observed in SEM micrographs. Our study thus suggests that mercury exposure caused stronger oxidative stress in shoots of P. vittata than in N. exaltata. Plants respond to oxidative stress by developing both enzymatic and non-enzymatic anti-oxidative systems (Apel and Hirt 2004). Within a cell, SOD is ubiquitous in all the subcellular locations. SOD constitutes the first line of defense against ROS by scavenging free radicals by means of catalyzing the dismutation of superoxide radicals in tissues (Alscher et al. 2002). In addition, CAT is an indispensable defensive anti-oxidative enzyme that removes excessive H2O2 by reducing H2O2 to H2O and hence protecting plant tissues from oxidative stress (Angela et al. 2004). In the present study, the only significant increases in the activities of SOD and CAT were observed in N. exaltata after mercury treatment, but not in P. vittata. This indicates that N. exaltata may remove ROS by DOS and CAT more rapidly than does P. vittata under mercury treatment. GR is responsible for regeneration of glutathione (GSH) by reducing oxidized glutathione (GSSG) in a NADPH-dependent reaction. GSH is not only an effective antioxidant during the GSH-ASC cycle (Ron 2002), but also a precursor for biosynthesis of phytochelatins (PCs). These PCs have been indicated to be induced in vivo as a possible detoxification mechanism for several heavy metals, such as cadmium, copper and arsenic (Gupta et al. 1998; Pickering et al. 2000; Zhu et al. 1999). Although GSH concentrations in both ferns increased under mercury exposure, GR activity in N. exaltata was significantly increased at the higher mercury level, while it did not change in P. vittata. These results suggest that N. exaltata is more tolerant to mercury stress since it takes up less Hg by roots and has less transport to shoots (even though statistic analysis did not show a significant difference among Hg concentrations) as well as it is more effective in establishing an enzymatic anti-oxidative system than P. vittata. The fact that P. vitatta has higher amounts of H2O2 and TBARS in shoots
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(Figs. 4, 5) is most likely a direct response to its higher mercury concentrations in shoots than those in N. exaltata (Table 1). However, enzymatic responses (SOD, CAT, and GR) significantly increased with Hg treatments in N. exaltata (Fig. 6) that actually had relatively low mercury concentrations in shoots (Table 2). This observation confirms that N. exaltata has a more effective anti-oxidant system than P. vittata. In line with this, Srivastava et al. (2005) also reported a similar conclusion for arsenic between tolerant and sensitive fern species. Ascorbate is another antioxidant involved in the nonenzymatic defense system against oxidative stress (Ron 2002). Besides the function of ASA as co-substrate of plant oxidase in the GSH-ASA cycle, it can directly react with superoxide, hydrogen peroxide, and hydroxyl radical or it can quench singlet oxygen directly (Singh et al. 2006). Our data show that N. exaltata regulated the ASA-DHA pool more efficiently than did P. vittata when exposed to mercury stress. ASA concentration was significantly increased in N. exaltata, but not always increased in P. vittata under mercury exposure (increased initially, then decreased). However, DHA concentration in P. vittata significantly increased with mercury exposure (Fig. 8a). The changes between ASA and DHA resulted in increases in the ratio of ASA/(ASA ? DHA) in N. exaltata, but decreases in P. vittata under mercury exposure. Similarly, Singh et al. (2006) reported that there were larger ratios of ASA/ (ASA ? DHA) in arsenic-tolerant fern species than in arsenic-sensitive species under arsenic stress. In addition, the lower ratios of ASA/DHA have also been observed in other sensitive plant species to various stresses, such as copper (Gupta et al. 1999) and heat (Dat et al. 1998). As an index of redox status in cells, the high ratio of ASA/ (ASA ? DHA) was found in meristem of the root cap of onions, which suggests that the ASA-DHA pool may play an important role in the regulation of plant growth by adjusting the redox status in cells (Co´rdoba-Pedregosa et al. 2003). However, the relationship between the regulation of redox status and plant resistance requires more study. In conclusion, mercury exposure triggered stronger oxidative stress (accumulation of H2O2 and TBARS) and phytotoxicity (leaf burning symptoms, break-down of thylakoid in chloroplast) in P. vittata than in N. exaltata. Although the roots of both ferns accumulated large amounts of mercury, N. exaltata showed less uptake of mercury by roots, less transport into shoots, as well as a more efficient anti-oxidative system than P. vittata under mercury stress. Mercury accumulation in the roots of these two ferns was much higher than in other plants reported in the literature. Neither fern may be suitable for phytoextraction because of their low transfer of mercury from roots to shoots, but N. exaltata may be useful for phytostabilization of soils or phytofiltration of mercury waste water since its roots took
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up a large amount of mercury without severe phytotoxic symptoms. Further studies are needed to completely understand the mechanisms of the mercury tolerance by N. exaltata and its potential in phytoremediation. Acknowledgments We thank Ms. Yunju Xia and Mr. Dean W. Patterson for chemical analyses. We also gratefully acknowledge the electronic microscopy study assistance provided by Ms. Amanda M. Lawrence. This research is supported by U.S. Department of Energy’s Office of Science and Technology through Cooperative Agreement DE-FC01-06EW-07040.
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