The morphogenic responses and phytochelatin
complexes by arsenic in Pteris vittata change in the
presence of cadmium
M. Ronzan
a,1, L. Zanella
a,1,2, L. Fattorini
a, F. Della Rovere
a, D. Urgast
b, S. Cantamessa
c,
A. Nigro
d, M. Barbieri
d, L. Sanità di Toppi
e, G. Berta
c, J. Feldmann
b, MM. Altamura
a,
G. Falasca
a,*
a Department of Environmental Biology, “Sapienza” University of Rome, Italy
b
Trace Element Speciation Laboratory, Department of Chemistry, University of Aberdeen, United Kingdom c
Dipartimento di Scienze e Innovazione Tecnologica dell’Università del Piemonte Orientale “A. Avogadro”, Alessandria, Italy d
Dipartimento di Scienze della Terra, “Sapienza” University of Rome, Italy e Department of Life Sciences, University of Parma, Italy
ABSTRACT
Arsenic (As) and cadmium (Cd) are toxic elements frequently present simultaneously in the environment. Pteris vittata L. (Chinese brake fern) is a natural As hyperaccumulator, able to accumulate very high levels of As in the fronds. The fern is also capable to adsorb Cd and to accumulate it, at moderate levels, in the root. To date, the mechanisms triggered by the fern in the presence of As and Cd are poorly known, and it is unknown whether Cd alters the fern As hyperaccumulating capabilities. The research aim was to analyse the responses of P. vittata when exposed simultaneously to As and Cd. In particular, in this work it was investigated whether Cd alters the capabilities/strategies that the fern activates to hyperaccumulate As. Results showed that co-exposure to Cd and As negatively affects P. vittata by inducing cyto- histological damage in the fronds, and reducing plant biomass. Cadmium presence increases As uptake but reduces As translocation to the frond. Co-exposure to Cd and As causes variations in the formation of As-phytochelatin complexes, mainly inducing the synthesis of long chain thiols binding As. Moreover, the co-exposure to Cd and As enhances extrusion of exudates containing As and Cd from the fronds. All together the data show that Cd alters the natural capabilities of the fern to hyperaccumulate As, but also suggest that P.vittata is able to contrast the toxicity due to As plus Cd, at least within defined limits.
The metalloid arsenic (As) and the metal cadmium (Cd) are widespread and persistent contaminants in soil and water. Inorganic As species, arsenate (As V) and arsenite (As III), and Cd2+ ions, are highly toxic for plants, humans and animals. Arsenate is a phosphate analogue and it can interfere with phosphate metabolism, whereas arsenite reacts with sulfhydryl groups of enzymes and proteins, leading to inhibition of cellular functions (Meharg and Hartley-Whitaker, 2002). Cadmium ions can cause several symptoms of toxicity such as leaf chlorosis, photosynthesis inhibition, alteration of water balance and oxida- tive stress (López-Millán et al., 2009; Aghaz et al., 2013; Yu et al., 2013). Cadmium and As are often present simultaneously in the environment (Groudev et al.,
2001; Kim et al., 2003), and can be readily absorbed by the plants, with negative effects on the growth. Moreover, the presence of
As and Cd in the environment affects the plant uptake of essential metals due to their chemical similarity with these elements, and to competition for the same cellular transporters/channels (Verbruggen et al., 2009). However, few studies are available on the combined effects of As and Cd on plant development, and on the detoxification strategies that specific plants can implement to contrast their toxicity.
The fern Pteris vittata L. is an As-hyperaccumulator, considered the best plant for phytoremediation of As-contaminated
environ-ments ( etMa al., 2001). Its growth characteristics, such as an easy and fast growth, large biomass, wide and extensive root system, as well as a wide geographic distribution, justify the successful use to reclaim As-contaminated sites (Danh et al., 2014 and references therein). The fern ability to absorb, and hyperaccumulate As is due to highly efficient defence mechanisms, either direct (i.e., by storing As in the frond vacuoles), or indirect (i.e., by activating antioxidant systems) (Lombi et al., 2002; Singh et al., 2006;
Verbruggen et al., 2009). In addition, proteomic analyses showed up-regulation of multiple forms of glyceraldehyde-3-phosphate
dehydrogenase, phosphoglycerate kinase, and enolase, suggesting a central role for glycolytic enzymes in arsenic metabolism and a putative arsenic transporter, PgPOR29, has been identified as an up-regulated protein by arsenic treatment (Bona et al., 2010).
The simultaneous presence of different, potentially toxic, elements gives rise to an amplification of the damage in the plants, but also induces them to increase the types, and the intensity, of the defence strategies. However, the mechanisms behind the tolerance to multiple contaminations are still widely unknown. This knowledge would be important to identify plant species able to activate these strategies/mechanisms and clean up polluted sites by more than an inorganic toxic element.
In the last decades, some reports have shown that P.vittata is also Cd-tolerant, although low levels of the metal induce damages
during the entire life cycle (Gupta and Devi 1994; Balestri et al.,2014). Furthermore, a limited information, and with contrasting results, is also available on the capability of P. vittata to accumulate and tolerate Cd and As, when simultaneously present (Xiao et al., 2008; Drava et al., 2012). In fact, Xiao et al. (2008) reported that a Cd-tolerant ecotype of P. vittata is able to extract effectively As and Cd
from the contaminated soil, and to accumulate not only As, but also Cd, in the fronds. On the contrary, when the fern was co-exposed to As and Cd in hydroponic culture, the Cd presence produced a stronger negative impact, reducing significantly the biomass and photosynthetic efficiency of the plant (Drava et al.,2012).
It is well known that plants exposed to toxic ions, including As and Cd, show an increase in low-molecular-weight thiols, such as cysteine,
g
-glutamylcysteine, glutathione (GSH) andphytochela-tins (PCs) (Grill et al., 1985; Cobbett, 2000; Sanità di Toppi et al.,2003; Srivastava et al., 2009). The SH group of these thiols, particularly of PC and of its precursor GSH, binds As, mainly As(III), and Cd, resulting in toxicity alleviation in numerous plants (Rea,2012 and references therein). This alleviation possibly occurs because As- or Cd–PC complexes can be transported and sequestrated into the vacuoles (Song et al.,
2010; Brunetti et al., 2015). In addition, in P. vittata fronds it has been shown that As induces an increase of GSH, suggesting
glutathionylation as a possible arsenic detoxification strategy (Singh et al., 2006; Bonaet al., 2011). The phytochelatins have the common structure (
g
-Glu-Cys)nGly, where n = 2-11, although PC2 and PC3 are the most common peptides (Cobbett and Goldsbrough, 2002). The ability to synthesize PCs depends on genetic factors and the availability of sulphate (Cobbett and Goldsbrough, 2002; Duan et al., 2012).
It has been reported that the PC levels in the root of some species, such as Arabidopsis thaliana and Oryza sativa, can affect the
translocation to aerial organs of toxic elements, i.e., As (Liu et al.,2010; Batista et al., 2014). Moreover, in the hyperaccumulator species, the PC synthesis is not always positively related with metal/metalloid tolerance (Ebbs et al., 2002; Zhao et al., 2003;Raab et al., 2004). In fact, in
P. vittata only very small amounts of As are complexed with PC and GSH in roots and/or fronds (Zhao et al.,2003; Raab et al., 2004; Bona et
al., 2011). The reduced formation of arsenite-PC-complexes in this fern has been reported to be one of the reasons for its efficient
root-to-frond As translocation (Su et al.,2008; Zhao et al., 2009). However, to date it is not known whether P. vittata is able to increase the
thiol-synthesis when exposed simultaneously to As and Cd, and if these complexes may have a role in the tolerance of more than one toxic element. The foliar extrusion of metals and/or metalloids is a detoxifica- tion strategy, however it seems to be restricted to few species, such as
Nicotiana tabacum (Choi et al., 2001; Zanella et al., 2016), Brassica juncea (Salt et al., 1995), Alyssum lesbiacum (Krämer et al., 1997), Arabidopsis halleri, and Vigna radiata (Küpper et al., 2000;Gupta and Bhatnagar 2015). Recently, P. vittata has been
demonstrated to be able to extrude As by specific frond structures, as a possible mechanism to tolerate high As doses (Cantamessa et al., 2015). However, it is unknown if the extrusion by the fronds also occurs under exposure to As and/or Cd.
The present research was aimed to evaluate the ability of this fern to accumulate and tolerate As and Cd, and to analyse the strategies carried out to overcome their toxicity. To the aim, P. vittata plants were exposed to Cd and/or As, and the metalloid/ metal levels and
peptide complexes, the cyto-histological mod- ifications in the frond, and Cd/As extrusion by the latter, were evaluated.
Our results show that the fern is able to counteract the combined metal and metalloid toxicity by strengthening the cyto-histological barriers in the frond, extruding Cd and As from the fronds, and inducing different As-PC complexes, and at different levels, in roots and fronds.
2. Materials and methods
2.1. Fern growth and metal/metalloid treatments
Spores of Pteris vittata L. were sterilized, hydrated and sown on Petri dishes containing salts of MS solution (Murashige and Skoog,1962), 20 g/l sucrose, and 8 g/l agar.
The spores germinated after 15 days after sowing. After other 15 days, the gametophytes were fully developed, and after other 30 days they developed into sporophytes. The sporophytes were separated from each other and transferred into sterile magenta containing quartz sand saturated with a solution of Hoagland salts (Hoagland and Arnon, 1938) at half concentration (i.e., Hoagland medium). The sporophytes were monthly transferred onto quartz sand saturated with refreshed Hoagland medium to allow their full development. One-year-old sporophytes were used in the experi- ments.
One-year-old sporophytes with about 4-cm-in-length fronds were transferred onto refreshed Hoagland medium supplemented or not (Control) with 100
m
M Na2HAsO .7H O (i.e., 100 As), 60m
MCdSO4 (i.e., 60 Cd) and 100
m
M Na2HAsO4.7H2O plus 60m
M CdSO4(i.e., 100 As + 60 Cd), and hydroponically cultured for 15 days. The pH of all media was adjusted to 5.8-6.0. All cultures were kept in a
controlled light and temperature regimes with a 16-h light/8 h dark photoperiod, an irradiance of 8.976
m
mol m—2s—1, a day/night temperature of 25/19 ○C, and 70% humidity. Ultra-pure water (Milli-Q) was used for all culture media.2.2. Morphological, cyto-histological and histochemical analyses
Ten sporophytes per treatment were weighed before the transfer into a new medium containing or not containing (Control) the metal and/or the metalloid, and were weighed again after 15 days to calculate the fresh weight increase.
Fronds randomly chosen from other 10 sporophytes per treatment were observed on the 15th day under a stereomicro- scope (Zeiss – Lumar. V12) to detect the necrotic areas. Two fronds from these sporophytes were fixed in 70% ethanol for the cyto- histological analyses. The cyto-histological analysis was carried out on the apical pinna because it is the site of a higher As
accumulation according to Sakai et al. (2010). The pinna was divided in three regions, i.e., apical, middle, and basal one. Samples from these regions were dehydrated, embedded in paraffin (melting point 52–54 ○C), cross-sectioned at 8
m
m with an automaticmicrotome (Microm HM 350 SV), and stained with
eosin and Carazzi’s haemalum. For the histochemical detection of lipids in the cell walls, sections from paraffin-embedded samples were stained with Sudan IV according to Czerednik et al. (2012). The cuticle thickness was evaluated on 100 epidermal cells coming from10 sections of the central part of the median region. Sections were stained with Sudan IV, and were randomly chosen among those of 5 fronds per treatment. Measurements were carried out using the Leica IM1000 image analysis software.
Unstained sections were observed under a ZEISS Axiolab HBO 50 microscope equipped with the specific filter sets (UV filters, EX BP 340–380 nm, LP 425 nm) suitable for lignin autofluorescence detection. All the histological and morphological images were acquired with a DC500 video camera applied to the microscope.
2.3. Elements determination
After 15 days of As and Cd exposure, 10 plants per treatment were taken up and thoroughly washed with deionised water. Fronds and roots were isolated and dried at 60 ○C for two days. Dried tissues were weighed and homogenized in an agata mortar. Afterwards, homogenized
material (250 mg) was treated with HNO3 (ultrapure 65%) and H2O2 (3:1) in glass flasks fitted with air- cooled condensers. Solutions were heated at 140 ○C for 3 h. After cooling the suspension was filtered in a 50-ml volumetric flask and diluted to the mark. Cd and As concentration
was measured by Inductively Coupled Plasma Mass Spectrometry (ICP-MS), using a Thermo Scientific X Series 2 spectrometer. This instrument operates in the helium collision cell mode to eliminate interference from isobaric polyatomic species via kinetic energy discrimination. The instrument was standardized with multi-element standards containing Cd and As at concentrations of approximately 0.1, 0.5, 1.0, and 10 ng/mL in matrix of 2% HNO3. Accuracy of elemental analysis was checked by carrying a standard reference material (SRM 1547 peach leaves, US NIST, Gaithersburg, MD) according to Tu and Ma (2005).
2.4. As-phytochelatin complex analyses
Plants from the different exposures were collected, divided in roots and fronds, put into liquid nitrogen and conserved at 80 ○C. When ready
to be analysed the plants were grinded with liquid nitrogen. For the As-phytochelatin extraction about 1 g of the plant material was weighed into a vial, in which 3 ml of 1% formic acid was added and left in ice for at least 15 min. The extracted solution was centrifuged at 13.000 rpm for 2 min. The supernatant was analysed with RP-HPLC coupled to ICP-MS/MS (8800 Agilent technologies) and ESI –MS (Orbitrap Thermo Scientific).
About 0.5 ml of the standards and samples were placed into the auto-sampler of the on-line coupled HPLC with ICP-MS and ESI – MS. The column used was Agilent Eclipse XDB-C18 (150 × 4.6 mm).
Volume injected 100
m
l; column was kept at 30○ C and autosamplercooled at 4○ C. Flow rate was 1 ml/min. The eluent A was 0.1% formic acid in water and eluent B 0.1% formic acid in methanol. The gradient
changed from 0 to 20% of eluent B from 0 to 20 min, then kept constant from 20 to 30 min at 20% eluent B and at 30 min decreased to 0% eluent B again.
The ICP-MS/MS was in organic mode, Pt-cones, PTA micro nebuliser, H2/O2 as reaction gas (1.1 l/min H2 and 0.3% O2) and Ge (10 ng ml—1) was the internal standard added post column via a T- piece. The following elements were monitored directly on the isotopic masses (Ge+ on m/z 72 and Cd+ on m/z 111) while As and S were monitored as their oxides (SO+ on m/z 48 and AsO+ on m/z 91). Quantification of the arsenic and sulphur species was done according to Bluemlein et al., 2008.
2.5. Microanalysis of frond exudates
After 15 days 10 pinnae of 10 fronds coming from 10 different plants per each treatment were examined under stereomicroscope (Zeiss – Lumar. V12) to detect possible crystal presence on the frond surface. The fronds were dried at 60 ○C for two days. Thereafter,
the samples were mounted on stubs and observed under an Environmental Scanning Electron Microscope (ESEM, Quanta 200, FEI Company, Eindhoven, The Netherlands). Energy Dispersive X-ray Spectroscopy (EDS) analysis with ZAF correction (EDAX Inc., Mahwah, USA) was performed to determine the quantitative amounts of elements.
2.6. Statistical analysis
Statistical analysis was performed using ANOVA test (one way) followed by Tukey’s post-test through GraphPad Prism 6.07 software. All the experiments were performed in triplicate with similar results.
3. Results
3.1. Co-exposure to cadmium and arsenic reduces P.vittata growth
A morphological analysis was carried out to assess whether/ how Cd affects the fern growth when combined to As for 15 days. The fronds of the fern cultured under the adopted in vitro conditions described in Materials and methods never differentiat- ed
sporangia. In the absence of the metalloid and of the metal (i.e., in the control treatment), the plants showed green fronds, with sporadic necrosis possibly as a consequence of natural senescence, and a developed root system (Fig. 1 A). The treatment with Cd alone and, mostly, the co-exposure of Cd and As, strongly reduced plant biomass, increased necrosis in the fronds, but enhanced the inception of new fronds in comparison with the control-treated plants. However, the new fronds were chlorotic and remained at the pastorale stage (Fig. 1 B and D, arrows). The As alone treatment did not reduce the plant growth and did not induce frond necrosis (Fig. 1C), as expected. The mean increase of plant fresh weight changed accordingly. In fact, the As treatment induced the highest growth, which was significantly higher than in the Control (P < 0.05), Cd alone (P < 0.01) and Cd plus As (P < 0.01) treatments. By contrast, in comparison with the controls, the fresh weight was significantly (P < 0.05) reduced for the co-exposed plants (Fig. 1 E).
3.2. Co-exposure to cadmium and arsenic induces cyto-histological damages in the frond
The histological analysis was carried out on the apical, median and basal part of the apical pinna of the frond coming from plants non treated (Control) or treated with 60 Cd, 100 As, and 100 As plus 60 Cd for 15 days.
The exposure to As alone did not cause any modification in the pinna tissue organization in comparison with the Control (Fig. 2A). In fact, in both cases the pinna showed dorsiventral structure, with a palisade parenchyma under the adaxial epidermis and a spongy parenchyma above the abaxial one and without differences related to the pinna region. Moreover, spongy parenchyma cells were lobed and separated by limited intercellular spaces (Fig. 2A), and the midrib exhibited a triangular/circular shape, with xylem and phloem surrounded by an endodermis with evident Casparian strips (Figs. 2 B–D and 3
Fig. 1. Phenotype of P. vittata non-treated (Control, A) and treated with either 60 mM CdSO4 (60 Cd, B) or 100 mM Na2HAsO4·7H2O (100 As, C) or with 100 mM
Na2HAsO4·7H2O plus 60 mM CdSO4 (100 As + 60 Cd, D), and plant mean fresh weight ( TSE) after 15 days of treatment (E). The arrows, in B and in Inset, show the fronds at
pastorale stage after 60 Cd and 100 As + 60 Cd treatments.
Letter a, P < 0.01 difference with respect to Cd and As + Cd treatments. Letter b, P < 0.05 difference with respect to the Control. Letter c, P < 0.05 with respect to the Control and Cd alone. Columns followed by the same letter are not significantly different. N = 10.
Fig. 2. Cross-sections of the basal (B, F, J), median (A, C, E,G, I, K) and apical (D, H, L) regions of the lamina (A, E, I) and midrib (B-D, F-H, J-L) of the apical pinna of the fern non treated (Control, A-D) or treated with 60 mM CdSO4 (60 Cd, E-H) or with 100 mM Na2HAsO4 7H2O plus 60 mM CdSO4 (100 As + 60 Cd, I-L) for 15 days. A — Lamina with lobed cells in the spongy tissue and with limited intracellular spaces. B-D, Midrib showing the vascular bundle surrounded by endodermis. E, Lamina showing wide intracellular
spaces. F, midrib with a very strong cell wall thickening in all tissues. G-H, hypertrophy in the parenchyma cells (G), crushing the endodermis (G-H, arrows in G), and cell wall thickening in the epidermis (G-H, and Inset in H). I, lamina showing an enhanced intercellular spaces in comparison with E. J-L, midrib with a strong cell wall thickening in all
tissues, hypertrophy in the parenchyma cells (J), collapse of the parenchyma cells, and crushing of the endodermis (K-L). Bars = 10 mm (Inset in H), 20 mm (G-I, K-L), 40 mm
(A-B, E-F, J) and 50 mm (D). A–B). In all regions, the adaxial epidermis, especially in correspondence of the midrib, showed a thin cuticle (Fig. 3E). On the contrary, the exposure to Cd alone deeply altered the pinna structure. In fact, a strong increase in the intercellular spaces of the lamina was observed under this treatment (Fig. 2E). In the basal region of the pinna, Cd caused a strong cell wall thickening in all the tissues around the midrib, including the epidermis (Figs. 2F, 3F–G), with that of the epidermal cells mainly due to lipid deposition (Fig. 3F-G), and that of the parenchyma cells to cellulose and lignin deposition (Fig. 3 H). In the median and apical regions of the pinna, Cd also induced hypertrophy in the parenchyma cells bordering the midrib (Fig. 2G-H), crushing the endodermis (Figs. 2 G–H, arrows, and 3 C). The external cell wall of the epidermis in the central-median region showed a mean thickness of 8.96( 0.5), 7.13( 0.4), and 4.11( 0.2) mm, in the Cd, As plus Cd and Control treatments, respectively. Thus, Cd induced a higher (P < 0.01) thickening in comparison with the other treatments, and this was due to an increased cuticle (Fig. 2G-, and Inset in H) as in the basal region (Fig. 3F–G).
The exposure to Cd plus As induced damages analogous, but more marked, to those caused by Cd alone both in the lamina and in the midrib (Figs. 2 I–L and 3 D), with the strongest damages again occurring in the median and apical regions. Moreover, metabolite deposition also characterized the cellular cell walls and the intercellular spaces of the midrib parenchyma (Fig. 3 I–J).
·
However, the epidermal cell wall thickening, even if significantly (P < 0.01) higher than in the Control, was reduced in comparison with Cd alone.
3.3. Cadmium increases As uptake but reduces its translocation to the frond
To evaluate whether the presence of Cd, combined with As, induced variations in As uptake and accumulation in the fern organs, an ICP-MS analysis was carried out on roots and fronds of plants exposed, or not, to 100 As, 60 Cd, and 100 As plus 60 Cd for 15 days. The control plants showed very negligible levels of As and Cd, but detectable due to traces of As and Cd in the growth media (Fig. 4 ). The roots of the plants treated with As alone showed As levels significantly (P < 0.01) lower than the roots of the plants exposed to As plus Cd (Fig. 4 ). On the contrary, the Cd accumulation in the root was significantly (P < 0.01) higher in the plants exposed to Cd alone in comparison with the plants co- exposed to As and Cd (Fig. 4 ). As expected, the highest As levels were found in the fronds in comparison with roots, however, the co-exposure to Cd and As significantly reduced As translocation to the fronds in comparison with As alone (Fig. 4 ). Cadmium was translocated to the fronds at very low levels, either in plants exposed to Cd alone or in plants exposed to As plus Cd, even if at a significant higher level (P < 0.05) in the latter ones (Fig. 4 ).
Fig. 3. Cross-sections of basal (A, C-D, E-H) and median (B, I-J) regions of the apical pinna at the midrib level in ferns non treated (Control, A-B, E) or treated with 60 mM CdSO4 (60 Cd, C, F-H) or with 100 mM Na2HAsO4 7H2O plus 60 mM CdSO4 (100 As + 60 Cd, D, I-J) for 15 days. A-D, midrib with endodermis showing regular (A-B), irregular
(C) and without (D) Casparian strips detected under UV light. E-G, Epidermis without lipid deposition (E) and with a lipid layer (F-G) after Sudan IV stain. H, lignin deposition in all cell walls shown by lignin autofluorescence. I-J, metabolite deposition in the intracellular spaces of the midrib parenchyma. Bars = 10 mm (E-H, J), 20 mm (A, C-D) and 40 mm (B, I).
Fig. 4. Mean concentrations (TSE) of As and Cd in roots and fronds of P.vittata not exposed (Control) and exposed for 15 days with either 100 mM Na2HAsO4 7H2O (100 As), or
60 mM CdSO4 (60 Cd), or with 100 mM Na2HAsO4 7H2O plus 60 mM CdSO4 (100 As + 60 Cd). Letters show statistical differences for the same element within the same organ;
Symbols indicate statistical differences for the same element, alone or combined, between different organs. a, P < 0.01 difference with 100 As. b, P < 0.01 difference with 100 As + 60 Cd. c, P < 0.01 difference with 100 As + 60 Cd. d, P < 0.05 difference with 60 Cd. *, P < 0.01 difference of As and Cd between root and frond. ○, P < 0.05 difference of As between root and frond. x, P < 0.01 difference of Cd between root and frond. Means of three replicates.
3.4. Cadmium induces variations in As-phytochelatin complexes in the fern exposed to As plus Cd
The fern exposed to As alone, and As plus Cd, was able to increase the synthesis of thiol compounds, such as GSH, and to induce the production of PCs. In both treatments, PCs, and GSH, forming complexes with As, were found (Figs. 5 and 6 and Supplementary information 1). In the roots and fronds of plants exposed to As alone, the presence of As and sulphur was detected in both organs (Fig. 5E-F and Supplementary information 1C-D). As-PC2, As-PC3 and GS-As-PC2 were the prevalent complexes, and As-PC3 in particular (Fig. 5A-D and Supplementary information 1A-B). These complexes were doubled in the roots than in the fronds (Fig. 5 A, C and B, D, in comparison, and Supplementary information 1A and B in comparison).
Also in roots and fronds of plants exposed to As plus Cd, the presence of As and sulphur was detected in both organs (Fig. 6G–H and Supplementary information 1G–H). The exposure to As plus Cd continued to induce the synthesis of As-PC2, As-PC3 (Fig. 6C–F) and GS-As-PC2 complexes (Supplementary information 1E-F), but also caused the appearance of the As-PC4 complex (Fig. 6 A–B). As- PC3 was the prevalent complex in both organs, but with a content five-fold higher in the fronds than the roots (Fig. 6C–D), contrary to what happened in the former organ when As alone was applied (Figs. 6 D and 5 B in comparison). In plants exposed to As with or without Cd the oxidized form of PC4, was also detected (fronds and roots, data not shown). In the control plants, the As-PCs complexes were never observed. No cadmium PC complexes were detected because they are labile in the extraction media and the mobile phase. These analytical conditions have however been used to stabilise the arsenic PC complexes (Bluemlein et al., 2009).
3.5. When combined with As, Cd induces the extrusion of As- and Cd- containing exudates from the frond
Stereomicroscope and scanning electron microscopy (SEM) analyses revealed the presence of exudates, either as saline secretions and discrete crystals, on the fern fronds cultured both in the absence and in the presence of As, Cd, and As plus Cd (Figs. 7 and 8 A,C,E,G ). The exudates were preferentially localized on the vasculature and on the edges of the pinnae (Fig. 7A-H). The exudates were more abundant on adaxial surface of the almost necrotic pinnae of the plants cultured in the presence of Cd, alone or combined with As (Fig. 7 C–E, and F–H, and Insets). To determine
the exudate element composition, an EDX analysis was performed for all treatments. The analysis showed that, independently of the treatment, the main constituents of exudates were C, O, and Ca, followed by K (Fig. 8B,D,F,H, and Inset). The exposure to As or Cd alone did not induce Cd or As extrusion (Fig. 8 D,F). By contrast, As and Cd were detected, as components of both saline secretions and crystals, when the fern was exposed to As plus Cd (Fig. 8H and Inset).
4. Discussion
The results showed that P. vittata biomass was significantly reduced by the exposure to both As plus Cd and Cd alone (Fig. 1). Visible symptoms of Cd toxicity in the fronds were observed (Figs. 2 and 3). These results are in contrast with previous data of Xiao et al. (2008)
showing that P.vittata is able to survive without damage in field, and in pot soil, containing Cd and As at concentrations higher than
those presently used. We exposed the fern to As plus Cd, or to Cd alone, through a hydroponic system. This system enables a complete bioavailability of the elements, as also shown by Drava et al. (2012), differently from the culture in soil used by Xiao et al. (2008). Thus, the difference between their and our results could be due to a higher bioavailability of the metal/metalloid caused by our culture conditions.
We observed that under the most stressful conditions for the fern, i.e., the exposure to Cd plus As, there was an increase in frond formation (Fig. 1D). This event can be related to changes in the morphogenetic responses that the plant carried out as a defence strategy to reduce the stress effects. It has been, in fact, reported that abiotic stresses inhibit cell elongation and induce localized cell divisions and alterations in cell differentiation that can determine the overproduction of adventitious organs such as roots and shoots
(Potters et al., 2007). In accordance, Cd exposure increases formation of lateral roots in Arabidopsis (Remans et al., 2012), and of
axillary culms in Miscanthus (Arduini et al., 2004).
Our results showed that P.vittata exposed to As alone produced various As-thiol complexes, both in roots and in fronds (Fig. 5) and, when the fern was co-exposed to As and Cd, the complexes increased in type, i.e., As-PC4 appeared, and in levels, i.e., the As- PC3 increased (Fig. 6). However, the increase in As-PC3 in the fronds was due to Cd presence, because a decrease in this complex was observed in the presence of As alone.
The levels of As-thiol complexes (As-PC3 in particular) were considerable, and higher in the roots than in the fronds, in contrast
Fig. 5. Identification by Orbitrap ESI–MS of As-PCs complexes and by ICP-MS of 32S and 75As in roots and fronds of P. vittata exposed to 100 mM Na2HAsO4 7H2O for 15 days.
A-B, As-PC3 levels in roots (A) and in fronds (B). C-D, As-PC2 levels in roots (C) and in fronds (D). E-F, 32S (gray line) and 75As (black line) levels in roots (E) and fronds (F).
with previous results in the same and other hyperaccumulating ferns (Zhao et al., 2003; Raab et al., 2004; Zhang et al., 2004;Vetterlein et
al., 2009), and in contrast with the only presence of As-PC2 complexes reported for P. vittata after As exposure (Zhaoet al., 2003; Bona
et al., 2011). These different results could be caused by the higher concentrations of arsenate, and its higher bioavailability due to the
hydroponic cultural system here used. The perception of high levels of As could trigger responses in the fern similar to those activated in the non-hyperaccumulator plants, e.g., the formation of As-thiol complexes at higher levels, as occurs in tobacco (Zanella et al., 2016).
Present data about As-PC4, As-PC3, As-PC2, and GS-As-PC2 under exposure to As plus Cd, in comparison with As alone (Figs. 6and 7), clearly show that Cd is very toxic for P. vittata. The thiols synthesis is related to the Cd level and its toxicity in numerous plants
(Cobbett, 2000; Sanità di Toppi et al., 2003). Thus, also in this fern the higher toxicity due to co-exposure to Cd and As can induce very high
Phytochelatins with longer chains are frequently related to the adaptative responses of the tolerant plants (Zhang et al., 2004). In this study, for the first time, we demonstrate that P.vittata is able to
Fig. 6. Identification by Orbitrap ESI–MS of As-PCs complexes and by ICP-MS of 32S and 75As in roots and fronds of P.vittata exposed to 100 mM Na2HAsO4 7H2O plus 60 mM
CdSO4 for 15 days. A-B, As-PC4 levels in roots (A) and in fronds (B). C-D, As-PC3 levels in roots (C) and in fronds (D). E-F, As-PC2 levels in roots (E) and in fronds (F). G-H, 32S (gray
line) and 75As (black line) levels in roots (G) and fronds (H).
Fig. 7. Stereomicroscope images of pinnae of fronds non exposed to As and/or Cd (Control, A), and exposed for 15 days with either 100 mM Na2HAsO4 7H2O (100 As, B), or 60
mM CdSO4 (60 Cd, C-E), or with 100 mM Na2HAsO4 7H2O plus 60 mM CdSO4 (100 As + 60 Cd, F-H) showing saline secretion and discrete crystals on the adaxial epidermis. Bars =
1 mm (A-D, F-H) and 0.5 mm (E and Insets in C, F and H).·
complex As with PC2 and GS-As-PC2, also in the frond. In addition, As also complexes with PC3 and PC4, both in the root than in frond, with the last two forms strongly enhanced and induced, respectively, by Cd. Thus, the role of PCs in the As tolerance of the fern does not seem to be limited, as previously reported (Zhao et al., 2003), but it appears relevant, and related to the toxicity levels that the fern has to support. Accordingly, in Helianthus annuus, Raab et al. (2007) have demonstrated that the formation of As–PC complexes is As concentration-dependent. Furthermore, higher levels of PCs are known to be positively related to the tolerance of As and/or Cd in Arabidopsis and tobacco
(Brunetti et al., 2011; Zanella et al., 2016). Years ago a cDNA for a phytochelatin synthase gene from P.vittata was cloned, and its expression
in Saccharomyces cerevisae conferred Cd resistance (Dong et al.,2005), suggesting that phytochelatins in P.vittata are functionally
similar to those of many other plants.
We observed that Cd levels were significantly higher in roots
than in fronds (Fig. 4 ) sustaining that P.vittata is able to absorb and accumulate Cd mostly in the root, whereas Cd transport to the frond
is reduced. These results are in accordance with those previously reported by Drava et al. (2012), using similar cultural conditions. On the contrary, Balestri et al. (2014) have reported that the fern is able to transport Cd to the fronds when the plant is exposed to Cd alone.
We showed that Cd presence significantly increased As uptake, but this was not associated with a greater transport to aerial organs (Fig. 4). This could be due to increased PCs levels in the root, that through the As-PCs complexes formation, and probably accumu- lation in the root cell vacuoles, allow a greater As influx, but not a bigger transport to the frond. It is known that As-PCs complexes formation in the roots reduces arsenite translocation from roots to shoots in Arabidopsis thaliana and in Oryza sativa (Batista et al.2014), and this has been
Numerous cyto-histological modifications in root and leaf have been described for plants exposed to toxic metals that are either the result of stress adaptative responses or the expression of damages suffered by the plant (Le Gall et al., 2015 and references therein). Arsenic alone also induces cytological modifications, e.g., in the root meristem of P. vittata (Forino et al., 2012). Our results show that cyto-histological alterations occur in the pinnae, but only when the fern was exposed to Cd plus As, or Cd alone (Figs. 2 and 3 ). The presence of Cd is a stressfull condition for the fern that required the development of different types of anatomical barriers, such as increased cell wall thickness, due to lignin over-deposition, and increased cutin deposition in the epidermal cells (Fig. 3). Moreover, also the crushing of the endodermal cells in the frond (Fig. 3C–D) could be attributed to the metal toxicity, similarly to what happens in the root endodermis of carrot exposed to Cd (Sanità di Toppi et al., 2012). The toxic elements reach the fronds through the xylem transport, thus the modifications here described can be interpreted as a fern strategy to limit the movement of toxic ions in the frond tissues. Recently it is reported that Cd induces morpho-physiological and proteomic changes in the leaves of sorghum, mostly determining cell wall modifications (Roy et al., 2016). The modifications may be enhanced by the level of metal toxicity, in fact, in our work, the simultaneous presence of As and Cd increased at least some of the modifications/alterations in the frond tissues. Additionally, the presence of metabolite deposition in the intercellular spaces and cell walls could be due to an increase in the synthesis of antioxidants, such as tocopherols. In fact, it has been reported that the antioxidant defences have an important role in metal stress response (Collin et al., 2008; Sunet al., 2010; Sanità di Toppi et al., 2012).
frond revealed the presence of both saline secretions and discrete crystals, and when the fern was exposed to As combined to Cd these extrusions contained As and Cd among the other elements
Fig. 8. SEM analysis on fern fronds showing saline secretion and crystals extrusion (A, C, E, G) and chemical composition of the excreted by EDS microanalysis (B, D, F, H). Ferns were not exposed (Control, A-B) or exposed to 100 mM Na2HAsO4 7H2O (100 As, C-D), or 60 mM CdSO4 (60 Cd, E-F), or with 100 mM Na2HAsO4 7H2O plus 60 mM CdSO4 (100 As
+ 60, Cd, G-H) for 15 days. Arsenic and cadmium presence in the extrusions, after exposure to As plus Cd, is shown by gray and black arrows, respectively (H). The Inset shows the concentrations (Conc.) of the elements (El.) expressed as atomic percentage after EDAX ZAF quantification (see Materials and methods).
(Figs. 7 and 8 ). Also the exudates of the fronds of the plants treated with higher Cd alone (120
m
M CdS04), and for longer periods (30 days), contained Cd (data not shown), and plants exposed to 334m
M arsenate (Na2HAsO4 7H2O) alone, for 60 days, are known to extrude crystals containing As (Cantamessa et al., 2015). Thus, the extrusion of Cd and/or As does not seem directly related to the combined presence of the two elements, but, rather, to the toxicitylevel. These results are in agreement with those reported in various other species after exposure to Cd and/or As or Zn, where they were extruded as crystals by the leaves through trichomes (Küpper et al., 2000; Choi et al., 2001; Gupta and Bhatnagar 2015; Zanella et al., 2016). In P.vittata, As has been demonstrated to be accumulated mostly in the epidermal cells of the pinnae (Lombi et al., 2002) and in the multicellular trichomes (Li et al., 2005). Moreover, it has been also reported that the fern is able to leach As from the frond (Yan et al., 2009), and, more recently, that the fern extrudes As through hydathodes-like structures (Cantamessa et al., 2015). Similar structures were never observed here, probably because of the slow growth caused by the in vitro culture, in fact in the present experiments generally
only trophophylls occurred. By contrast, hydathodes-like structures have been observed by Cantamessa et al. (2015) in the trophosporophylls, i.e., fronds with trophic and reproductive functions. Thus, it is possible that integrate mecha- nisms to extrude toxic elements are activated by the fern in relation to its physiological stage, cultural conditions and metal toxicity.
5. Conclusions
In the present work we demonstrate that co-exposure to Cd and As affects the natural capabilities of Pteris vittata to
hyper-accumulate As. Cadmium reduces fern biomass, changes the morphological responses and thiolic metabolism induced by As, causes cyto-histological damage in the fronds, increases As uptake, but restricts As translocation to the frond, and enhances extrusion of exudates containing As and Cd from the frond. Cadmium alters the As hyperaccumulation capability of the fern. However, the fern is able to contrast, within defined limits, the toxicity due to Cd combined to As, highlighting its possible use in phytoremediation programs of soils where the two toxic elements are present at moderate concentrations.
Conflict of interest
The authors declare that they have no conflict of interest.
Author contributions
RM and ZL designed and carried out the research. FL and DRF contributed to carry out the histological analyses. UD and FJ carried out the thiol-As complex analyses. CS and BG carried out the SEM- EDS analyses on frond crystals. NA and BM carried out the ICP analyses. SdiTL, AMM and GF analysed data and wrote the manuscript. All authors read and approved the manuscript.
This work was supported by Progetti Ateneo Sapienza University of Rome (Year 2013 — prot. C26A138JM3 and Year 2014 — prot. C26A14AFZ7) to G.F.
Acknowledgements
The authors are grateful to Prof. L. Cornara from Genoa University for providing P. vittata L. spores.
References
Aghaz, M., Bandehagh, A., Aghazade, E., Toorchi, M., Ghassemi-Gholezani, K., 2013. Effects of Cadmium stress on some growth and physiological characteristics in Dill ( Anethum graveolens ) ecotypes. Int. J. Agric.: Res Rev. 3, 409 – 413 .
Arduini, I., Masoni, A., Mariotti, M., Ercoli, L., 2004. Low cadmium application increase miscanthus growth and cadmium translocation. Environ. Exp. Bot. 52, 89 – 100. doi:http://dx.doi.org/10.1016/j.envexpbot.2004.01.001.
Balestri, M., Ceccarini, A., Forino, L.M.C., Zelko, I., Martinka, M., Lux, A., Ruf fi ni Castiglione, M., 2014. Cadmium uptake, localization and stress-induced morphogenic response in the fern Pteris vittata . Planta 239, 1055 – 1064. doi: http://dx.doi.org/10.1007/s00425-014-2036-z.
Batista, B.L., Nigar, M., Mestrot, A., Rocha, B.A., Barbosa Júnior, F., Price, A.H., Raab, A., Feldmann, J., 2014. Identification and quantification of phytochelatins in roots of rice to long-term exposure: evidence of individual role on arsenic accumulation and translocation. J. Exp. Bot. 65, 1467 – 1479. doi:http://dx.doi.org/10.1093/jxb/ eru018.
Bluemlein, K., Raab, A., Meharg, A.A., Charnock, J.M., Feldmann, J., 2008. Can we trust mass spectrometry for determination of arsenic peptides in plants: comparison of LC-ICP-MS and LC-ES-MS/LC-ICP-MS with XANES/EXAFS in analysis of Thunbergia alata . Anal. Bioanal. Chem. 390, 1739 – 1751. doi:http://dx.doi.org/ 10.1007/s00216-007-1724-y. Bluemlein, K., Raab, A., Feldmann, J., 2009. Stability of arsenic peptides in plant extracts: off-line versus on-line parallel elemental and molecular mass spectrometric
detection for liquid chromatographic separation. Anal. Bioanal. Chem. 393, 357 – 436. doi:http://dx.doi.org/10.1007/s00216-008-2395-z.
Bona, E., Cattaneo, C., Cesaro, P., Marsano, F., Lingua, G., Cavaletto, M., Berta, G., 2010. Proteomic analysis of Pteris vittata fronds: two arbuscular mycorrhizal fungi differentially modulate protein expression under arsenic contamination.
Proteomics 10, 3811 – 3834. doi:http://dx.doi.org/10.1002/pmic.200900436.
Bona, E., Marsano, F., Massa, N., Cattaneo, C., Cesaro, P., Argese, E., Sanità di Toppi, L., Cavaletto, M., Berta, G., 2011. Proteomic analysis as a tool for investigating arsenic stress in Pteris vittata roots colonized or not by arbuscular mycorrhizal symbiosis. J. Prot. 74, 1338 – 1350. doi:http://dx.doi.org/10.1016/j. jprot.2011.03.027.
Brunetti, P., Zanella, L., Proia, A., De Paolis, A., Falasca, G., Altamura, M.M., Sanita ‘ di Toppi, L., Costantino, P., Cardarelli, M., 2011. Cadmium tolerance and phytochelatin content of Arabidopsis seedlings over-expressing the phytochelatin synthase gene AtPCS1 . J. Exp. Bot. 62, 5509 – 5519. doi:http://dx. doi.org/10.1093/jxb/err228.
Brunetti, P., Zanella, L., De Paolis, A., Di Litta, D., Cecchetti, V., Falasca, G., Barbieri, M., Altamura, M.M., Costantino, P., Cardarelli, M., 2015. Cadmium-inducible expression of the ABC-type transporter AtABCC3 increases phytochelatin- mediated cadmium tolerance in Arabidopsis . J. Exp. Bot. 66, 3815 – 3829. doi: http://dx.doi.org/10.1093/jxb/erv185.
Cantamessa, S., D ’ Agostino, G., Berta, G., 2015. Hydathode structure and localization in Pteris vittata fronds and evidence for their involvement in arsenic leaching. Plant Biosyst. doi:http://dx.doi.org/10.1080/11263504.2015.1012135.
Choi, Y.E., Harada, E., Wada, M., Tsuboi, H., Morita, Y., Kusano, T., Sano, H., 2001. Detoxification of cadmium in tobacco plants: formation and active excretion of crystals containing cadmium and calcium through trichomes. Planta 213, 45 – 50. doi:http://dx.doi.org/10.1007/s004250000487.
Cobbett, C., Goldsbrough, P., 2002. Phytochelatins and metallothioneins: roles in heavy metal detoxification and homeostasis. Annu. Rev. Plant Biol. 53, 159 – 182. doi:http://dx.doi.org/10.1146/annurev.arplant.53.100301.135154.
Cobbett, C.S., 2000. Phytochelatins and their roles in heavy metal detoxification.
Plant Physiol. 123, 825 – 832. doi:http://dx.doi.org/10.1104/pp.123.3.825.
Collin, V.C., Eymery, F., Genty, B., Rey, P., Havaux, M., 2008. Vitamin E is essential for the tolerance of Arabidopsis thaliana to metal-induced oxidative stress. Plant Cell Environ. 31, 244 – 257. doi:http://dx.doi.org/10.1111/j.1365- 3040.2007.01755.x.
Czerednik, A., Busscher, M., Bielen, B.A.M., Wolters-Arts, M., de Maagd, R.A., Angenent, G.C., 2012. Regulation of tomato fruit pericarp development by an interplay between CDKB and CDKA1 cell cycle genes. J. Exp. Bot. 63, 2605 – 2617. doi:http://dx.doi.org/10.1093/jxb/err451.
Danh, L.T., Truong, P., Mammucari, R., Foster, N., 2014. A critical review of the arsenic uptake mechanisms and phytoremediation potential of Pteris vittata . Int. J. Phytoremediat. 16, 429 – 453. doi:http://dx.doi.org/10.1080/
15226514.2013.798613.
Dong, R., Formentin, E., Losseso, C., Carimi, F., Benedetti, P., Terzi, M., Lo Schiavo, F., 2005. Molecular cloning and characterization of a phytochelatin synthase gene, PvPCS1 , from Pteris vittata L. J. Ind. Microbiol. Biotechnol. 32, 527 – 533. doi: http://dx.doi.org/10.1007/s10295-005-0234-1.
Drava, G., Roccotiello, E., Minganti, V., Manfredi, A., Cornara, L., 2012. Effects of cadmium and arsenic on Pteris vittata under hydroponic conditions. Environ. Toxicol. Chem. 31, 1375 – 1380. doi:http://dx.doi.org/10.1002/etc.1835.
Duan, G., Kamiya, T., Ishikawa, S., Arao, T., Fujiwara, T., 2012. Expressing ScACR3 in rice enhanced arsenite efflux and reduced arsenic accumulation in rice grains. Plant Cell Physiol. 53, 154 – 163. doi:http://dx.doi.org/10.1093/pcp/pcr161.
Ebbs, S., Lau, I., Ahner, B., Kochian, L., 2002. Phytochetatin synthesis is not responsible for Cd tolerance in the Zn/Cd hyperaccumulator Thlaspi caerulescens(J. & C.Presl). Planta 214, 635 – 640. doi:http://dx.doi.org/10.1007/ s004250100650.
Forino, L.M.C., Ruffini Castiglione, M., Bartoli, G., Balestri, M., Andreucci, A., Tagliasacchi, A.M., 2012. Arsenic-induced morphogenic response in roots of arsenic hyperaccumulator fern Pteris vittata . J. Hazard. Mater. 235-236, 271 – 278. doi:http://dx.doi.org/10.1016/j.jhazmat.2012.07.051.
Grill, E., Winnacker, E.L., Zenk, M.H., 1985. Phytochelatins: the principal heavy- metal complexing peptides of higher plants. Science 230, 674 – 676. doi:http:// dx.doi.org/10.1126/science.230.4726.674.
Groudev, S.N., Spasova, I.I., Georgiev, P.S., 2001. In situ bioremediation of soils contaminated with radioactive elements and toxic heavy metals. Int. J. Miner. Process. 62, 301 – 308. doi:http://dx.doi.org/10.1016/S0301-7516(00)00061-2.
Gupta, P., Bhatnagar, A.K., 2015. Spatial distribution of arsenic in different leaf tissues and its effect on structure and development of stomata and trichomes in mung bean Vigna radiata (L.) Wilczek. Environ. Exp. Bot. 109, 12 – 22. doi:http:// dx.doi.org/10.1016/j.envexpbot.2014.08.001.
Gupta, M., Devi, S., 1994. Chronic toxicity of cadmium in Pteris vittata , a roadside fern. Ecotoxicology 3, 235 – 247. doi:http://dx.doi.org/10.1007/BF00117990.
Hoagland, D.R., Arnon, D.I., 1938. The Water-culture Method for Growing Plants Without Soil. Berkeley, California, University of California, College of Agriculture, Agricultural Experiment Station.
Küpper, H., Lombi, E., Zhao, F.J., McGrath, S.P., 2000. Cellular compartmentation of cadmium and zinc in relation to other elements in the hyperaccumulator Arabidopsis halleri . Planta 212, 75 – 84. doi:http://dx.doi.org/10.1007/ s004250000366.
Kim, M.J., Ahn, K.H., Jung, Y., Lee, S., Lim, B.R., 2003. Arsenic, cadmium, chromium, copper, lead, and zinc contamination in mine tailings and nearby streams of three abandoned mines from Korea. Bull. Environ. Contam. Toxicol. 70, 942 – 947. doi:http://dx.doi.org/10.1007/s00128-003-0073-6.
Krämer, U., Grime, G.W., Smith, J.A.C., Hawes, C.R., Baker, A.J.M., 1997. Micro-PIXE as a technique for studying nickel localization in leaves of the hyperaccumulator plant Alyssum lesbiacum . Nucl. Inst. Methods Phys. Res. B 130, 346 – 350. doi: http://dx.doi.org/10.1016/S0168-583X(97)00368-6.
López-Millán, A.F., Sagardoy, R., Solanas, M., Abadía, A., Abadía, J., 2009. Cadmium toxicity in tomato ( Lycopersicon esculentum ) plants grown in hydroponics.
Environ. Exp. Bot. 65, 376 – 385. doi:http://dx.doi.org/10.1016/j. envexpbot.2008.11.010.
Le
Gall, H., Philippe, F., Domon, J.M., Gillet, F., Pelloux, J., Rayon, C., 2015. Cell wall metabolism in response to abiotic stress. Plants 4, 112 – 166. doi:http://dx.doi. org/10.3390/plants4010112.
Li, W., Chen, T., Chen, Y., Lei, M., 2005. Role of trichome of Pteris vittata L. in arsenic hyperaccumulation. Sci. China Ser. C 48, 148 – 154. doi:http://dx.doi.org/10.1007/ BF02879667.
Liu, W.J., Wood, B.A., Raab, A., McGrath, S.P., Zhao, F.J., Feldmann, J., 2010.
Complexation of arsenite with phytochelatins reduces arsenite efflux and translocation from roots to shoots in Arabidopsis . Plant Physiol. 152, 2211 – 2221. doi:http://dx.doi.org/10.1104/pp.109.150862.
Lombi, E., Zhao, F.J., Fuhrmann, M., Ma, L.Q., McGrath, S.P., 2002. Arsenic distribution and speciation in the fronds of the hyperaccumulator Pteris vittata . New Phytol. 156, 195 – 203. doi:http://dx.doi.org/10.1046/j.1469-8137.2002.00512.x.
Ma, L.Q., Komar, K.M., Tu, C., Zhang, W., Cai, Y., Kennelley, E.D., 2001. A fern that hyperaccumulates arsenic. Nature 409, 579. doi:http://dx.doi.org/10.1038/
35054664.
Meharg, A.A., Hartley-Whitaker, J., 2002. Arsenic uptake and metabolism in arsenic resistant and nonresistant plant species. New Phytol. 154, 29 – 43. doi:http://dx. doi.org/10.1046/j.1469-8137.2002.00363.x.
Murashige, T., Skoog, F., 1962. A revised medium for rapid growth and bio assays with tobacco tissue cultures. Physiol. Plant. 15, 473 – 497. doi:http://dx.doi.org/ 10.1111/j.1399-3054.1962.tb08052.x.
Potters, G., Pasternak, T.P., Guisez, Y., Palme, K.J., Jansen, M.A.K., 2007. Stress- induced morphogenic responses: growing out of trouble? Trends Plant Sci. 12, 98 – 105. doi:http://dx.doi.org/10.1016/j.tplants.2007.01.004.
Raab, A., Feldmann, J., Meharg, A.A., 2004. The nature of arsenic-phytochelatin complexes in Holcus lanatus and Pteris cretica . Plant Physiol. 134, 1113 – 1122. doi: http://dx.doi.org/10.1104/pp.103.033506.
Raab, A., Ferreira, K., Meharg, A.A., Feldmann, J., 2007. Can arsenic-phytochelatin complex formation be used as an indicator for toxicity in Helianthus annuus ? J. Exp. Bot. 58, 1333 – 1338. doi:http://dx.doi.org/10.1093/jxb/erl300.
Rea, P.A., 2012. Phytochelatin synthase: of a protease a peptide polymerase made.
Physiol. Plant. 145, 154 – 164. doi:http://dx.doi.org/10.1111/j.1399- 3054.2012.01571.x.
Remans, T., Thijs, S., Truyens, S., Weyens, N., Schellingen, K., Keunen, E., Gielen, H., Cuypers, A., Vangronsveld, J., 2012. Understanding the development of roots exposed to contaminants and the potential of plant-associated bacteria for
optimization of growth. Ann. Bot. 110, 239 – 252. doi:http://dx.doi.org/10.1093/ aob/mcs105.
Roy, S.K., Cho, S.W., Kwon, S.J., Kamal, A.H.M., Kim, S.W., Oh, M.W., Lee, M.S., Chung,
K.Y., Xin, Z., Woo, S.H., 2016. Morpho-physiological and proteome level responses to cadmium stress in sorghum. PLoS One 11, e0150431. doi:http://dx. doi.org/10.1371/journal.pone.0150431.
Sakai, Y., Watanabe, T., Wasaki, J., Senoura, T., Shinano, T., Osaki, M., 2010. Influence of arsenic stress on synthesis and localization of low-molecular-weight thiols inPteris vittata . Environ. Pollut. 158, 3663 – 3669. doi:http://dx.doi.org/10.1016/j. envpol.2010.07.043.
Salt, D.E., Prince, R.C., Pickering, I.J., Raskin, I., 1995. Mechanisms of cadmium mobility and accumulation in Indian mustard. Plant Physiol. 109, 1427 – 1433. doi:http://dx.doi.org/10.1104/pp.109.4.1427.
Sanità di T oppi, L., Gremigni, P . , P a wli k -Skow r o n ´ ska, B., Prasad, M.N.V., Cobbett, C.S.,
2003. Response to heavy metals in plants: a molecular approach. In: Sanità di T oppi, L., P a wli k -Skow r o n ´ ska, B. (Eds.), Abiotic S tresses in Plants. Kluwer Academic Publishers, Dordrecht, The Netherlands, pp. 133 – 156. doi:http://dx. doi.org/10.1007/978-94-017-0255-3_6.
Sanità di Toppi, L., Vurro, E., De Benedictis, M., Falasca, G., Zanella, L., Musetti, R., Lenucci, M.S., Dalessandro, G., Altamura, M.M., 2012. A bifasic response to cadmium stress in carrot: early acclimatory mechanisms give way to root collapse further to prolonged metal exposure. Plant Physiol. Bioch. 58, 269 – 279. doi:http://dx.doi.org/10.1016/j.plaphy.2012.07.002.
Singh, N., Ma, L.Q., Srivastava, M., Rathinasabapathi, B., 2006. Metabolic adaptations to arsenic-induced oxidative stress in Pteris vittata L and Pteris ensiformis L. Plant Sci. 170, 274 – 282. doi:http://dx.doi.org/10.1016/j.plantsci.2005.08.013.
Song, W.Y., Park, J., Mendoza-Cózatl, D.G., Suter-Grotemeyer, M., Shim, D., Hörtensteiner, S., Geisler, M., Weder, B., Rea, P.A., Rentsch, D., Schroeder, J.I., Lee,
Y.,
Martinoia, E., 2010. Arsenic tolerance in Arabidopsis is mediated by two ABCC- type phytochelatin transporters. Proc. Natl. Acad. Sci. USA 107, 21187 – 21192. doi:http://dx.doi.org/10.1073/pnas.1013964107.
Srivastava, S., Srivastava, A.K., Suprasanna, P., D ’ Souza, S.F., 2009. Comparative biochemical and transcriptional profiling of two contrasting varieties of Brassicajuncea L. in response to arsenic exposure reveals mechanisms of stress perception and tolerance. J. Exp. Bot. 60, 3419 – 3431. doi:http://dx.doi.org/ 10.1093/jxb/erp181.
Su,
Y.H., McGrath, S.P., Zhu, Y.G., Zhao, F.J., 2008. Highly ef ficient xylem transport of arsenite in the arsenic hyperaccumulator Pteris vittata . New Phytol. 180, 434 – 441. doi:http://dx.doi.org/10.1111/j.1469-8137.2008.02584.x.
Sun, X., Zhang, J., Zhang, H., Ni, Y., Zhang, Q., Chen, J., Guan, Y., 2010. The responses of Arabidopsis thaliana to cadmium exposure explored via metabolite profiling. Chemosphere 78, 840 – 845. doi:http://dx.doi.org/10.1016/j. chemosphere.2009.11.045.
Tu, C., Ma, L.Q., 2005. Effects of arsenic on concentration and distribution of nutrients in the fronds of the arsenic hyperaccumulator Pteris vittata L. Environ. Pollut. 135, 333 – 340. doi:http://dx.doi.org/10.1016/j.envpol.2004.03.026.
Verbruggen, N., Hermans, C., Schat, H., 2009. Mechanisms to cope with arsenic or cadmium excess in plants. Curr. Opin. Plant Biol. 12, 364 – 372. doi:http://dx.doi. org/10.1016/j.pbi.2009.05.001.
Vetterlein, D., Wesenberg, D., Nathan, P., Bräutigam, A., Schierhorn, A., Mattusch, J., Jahn, R., 2009. Pteris vittata -revisited: uptake of As and its speciation, impact of P, role of phytochelatins and S. Environ. Pollut. 157, 3016 – 3024. doi:http://dx. doi.org/10.1016/j.envpol.2009.05.057.
Xiao, X., Chen, T., An, Z., Lei, M., Huang, Z., Liao, X., Liu, Y., 2008. Potential of Pterisvittata L. for phytoremediation of sites co-contaminated with cadmium and arsenic: the tolerance and accumulation. J. Environ. Sci. 20, 62 – 67. doi:http://dx. doi.org/10.1016/S1001-0742(08)60009-1.
Yan, X.L., Liao, X.Y., Chen, T.b., 2009. Leaching potential of arsenic from Pteris vittata
L. under field conditions. Sci. Total Environ. 408, 425 – 430;1;. doi:http://dx.doi. org/10.1016/j.scitotenv.2009.09.035.
Yu, L., Gao, R., Shi, Q., Wang, X., Wei, M., Yang, F., 2013. Exogenous application of sodium nitroprusside alleviated cadmium induced chlorosis, photosynthesis inhibition and oxidative stress in cucumber. Pakistan J. Bot. 45, 813 – 819 .
Zanella, L., Fattorini, L., Brunetti, P., Roccotiello, E., Cornara, L., D ’ Angeli, S., Della Rovere, F., Cardarelli, M., Barbieri, M., Sanità di Toppi, L., Degola, F., Lindberg, S., Altamura, M.M., Falasca, G., 2016. Overexpression of AtPCS1 in tobacco increases arsenic and arsenic plus cadmium accumulation and detoxification. Planta 243, 605 – 622. doi:http://dx.doi.org/10.1007/s00425-015-2428-8.
Zhang, W., Cai, Y., Downum, K.R., Ma, L.Q., 2004. Thiol synthesis and arsenic hyperaccumulation in Pteris vittata (Chinese brake fern). Environ. Pollut. 131, 337 – 345. doi:http://dx.doi.org/10.1016/j.envpol.2004.03.010.
Zhao, F.J., Wang, J.R., Barker, J.H.A., Schat, H., Bleeker, P.M., McGrath, S.P., 2003. The role of phytochelatins in arsenic tolerance in the hyperaccumulator Pterisvittata . New Phytol. 159, 403 – 410. doi:http://dx.doi.org/10.1046/j.1469- 8137.2003.00784.x.
