Mercury mobility and effects in the salt-marsh plant

Science of the Total Environment 650 (2019) 111–120

Contents lists available at ScienceDirect

Science of the Total Environment journal homepage: www.elsevier.com/locate/scitotenv

Mercury mobility and effects in the salt-marsh plant Halimione portulacoides: Uptake, transport, and toxicity and tolerance mechanisms Maria Teresa Cabrita a,⁎,1, Bernardo Duarte b, Rute Cesário a,c, Ricardo Mendes a, Holger Hintelmann d, Kevin Eckey d,e, Brian Dimock d, Isabel Caçador b, João Canário c a

Instituto do Mar e da Atmosfera (IPMA), Rua Alfredo Magalhães Ramalho, 6, 1495-006 Algés, Lisboa, Portugal MARE — Marine and Environmental Sciences Centre, Faculty of Sciences, University of Lisbon, Campo Grande, 1749-016 Lisbon, Portugal Centro de Química Estrutural, Instituto Superior Técnico, Universidade de Lisboa, Av. Rovisco Pais, Lisboa 1, 1049-001 Lisboa, Portugal d Water Quality Centre, Trent University, 1600 West Bank Drive, Peterborough, ON K9J 0G2, Canada e Institute of Inorganic and Analytical Chemistry, University of Muenster, Schlossplatz 2, 48149 Munster, Germany b c

H I G H L I G H T S

G R A P H I C A L

A B S T R A C T

• Mercury mobility in H. portulacoides assessed by stable isotope enriched Hg and hydroponics. • Mercury (THg, MMHg) is mainly accumulated into the roots of H. portulacoides. • Direct translocation of THg and MMHg occurs from roots to aerial parts of the plant. • Temperature and PAR influenced Hg content in stems and roots pointing to Hg release. • Low levels of Hg can impact H. portulacoides photochemistry with prolonged exposure.

a r t i c l e

i n f o

Article history: Received 8 April 2018 Received in revised form 23 August 2018 Accepted 24 August 2018 Available online 28 August 2018 Editor: D. Barcelo Keywords: Mercury Methylmercury Halimione portulacoides Accumulation Translocation Photochemical responses Salt marshes

a b s t r a c t The plant Halimione portulacoides, an abundant species widely distributed in temperate salt-marshes, has been previously assessed as bioindicator and biomonitor of mercury contamination in these ecosystems. The present study aims to assess uptake and distribution of total mercury (THg) and methylmercury (MMHg) within H. portulacoides, potential mercury release by volatilization through leaves, and toxicity and tolerance mechanisms by investigating plant photochemical responses. Stem cuttings of H. portulacoides were collected from a salt-marsh within the Tagus estuary natural protected area, and grown under hydroponic conditions. After root development, plants were exposed to 199HgCl2 and CH201 3 HgCl, and sampled at specific times (0, 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days)). After exposure, roots, stems and leaves were analysed for total 199 Hg (T199Hg) and MM201Hg content. Photobiology parameters, namely efficiency and photoprotection capacity, were measured in leaves. Both THg and MMHg were incorporated into the plant root system, stems and leaves, with roots showing much higher levels of both isotope enriched spikes than the other plant tissues. Presence of both mercury isotopes in the stems and leaves and high significant correlations found between roots and stems, and stems and leaves, for both THg and MMHg concentrations, indicate Hg translocation between the roots and above-ground organs. Long-term uptake in stems and leaves, leading to higher Hg content, was more influenced by temperature and radiation than short-term uptake. However, the relatively low levels of

⁎ Corresponding author at: CEG/IGOT, University of Lisbon, Rua Branca Edmée Marques, 1600-276 Lisbon, Portugal. E-mail address: [email protected] (M.T. Cabrita). 1 Present affiliation: Centro de Estudos Geográficos (CEG), Instituto de Geografia e Ordenamento do Território (IGOT), University of Lisbon, Rua Branca Edmée Marques, 1600-276 Lisbon, Portugal.

https://doi.org/10.1016/j.scitotenv.2018.08.335 0048-9697/© 2018 Elsevier B.V. All rights reserved.

112

M.T. Cabrita et al. / Science of the Total Environment 650 (2019) 111–120

both THg and MMHg in the aerial parts of the plant, which were influenced by temperature and radiation, support the possibility of mercury release by stems and leaves, probably via stomata aperture, as a way to eliminate toxic mercury. Regarding photochemical responses, few differences between control and exposed plants were observed, indicating high tolerance of this salt marsh plant to THg and MMHg. © 2018 Elsevier B.V. All rights reserved.

1. Introduction Mercury (Hg) is one of the most dangerous anthropogenic pollutants because of its high toxicity (Driscoll et al., 2013), and a major threat to coastal ecosystems mostly due to accumulation and biomagnification through food webs (Gworek et al., 2016). Mercury exists in many forms, and the most toxic ones are the organic mercury compounds, in particular methylmercury (MMHg), due to its high solubility in lipids, enhancing the potential for biological uptake and bioconcentration (Clayden et al., 2013). Methylmercury is made available by transformation of inorganic mercury (Hg(II)) through a complex set of biologically mediated chemical reactions under anaerobic conditions (Gworek et al., 2016; Randall et al., 2013). This organic form is a potent neurotoxin with harmful effects on reproduction and neural development in fish and mammals, representing most of the mercury content in contaminated fish and human bodies (Burgess and Meyer, 2008; Hong et al., 2012). Although anthropogenic Hg emissions have decreased by half over the last decades (Obrist et al., 2018; Pacyna et al., 2001), mercury contamination is still a global issue due to the long-range transport of this persistent pollutant across several environmental compartments (Fitzgerald and Lamborg, 2003; Sonke et al., 2013). Biological mercury hotspots found worldwide are a major concern to human populations and the ecosystems on which they depend (Driscoll et al., 2013; Evers et al., 2007; Obrist et al., 2018). Salt marshes are among Earth's most productive (Barbier et al., 2011; Giblin and Weider, 1992; Odum, 1971) and highly relevant ecosystems supporting vital ecosystem functions (Costanza et al., 1997), such as primary productivity, nutrient, metal and metalloid biogeochemical recycling, wildlife habitat, and shoreline stabilization (Mitsch and Gosselink, 2000). When located near polluted areas, these ecosystems are receivers of municipal, industrial and agricultural waste (Anjum et al., 2014a; Caçador et al., 2009; Gupta and Chandra, 1998). Although the global estimates of 45,000 to 54,951 km2 of salt marsh area (Greenberg et al., 2006; Mcowen et al., 2017; Pendleton et al., 2012) only correspond to about 0.07% of the total land surface, some salt marshes can act as sinks for metals and metalloids (Caçador et al., 1993, 1996; Hung and Chmura, 2006; Weis and Weis, 2004; Williams et al., 1994). Mercury inputs to waste are generally linked to effluents from metal production, pulp industries and chloralkali plants (Lindqvist et al., 1991). Whilst salt marshes can have an important role in reducing contamination of adjoining areas (Jacob and Otte, 2003), impairment of this function may occur as levels of contaminants rise. Once released into salt marshes, mercury can interact with both sediments and pore waters (Canário et al., 2005), and become available to the biota (Suntornvongsagul et al., 2007). Halophytes such as Halimione portulacoides, Sarcocornia sp. and Spartina maritima, which colonize and produce elevated biomass in salt marshes, retain metals/metaloids mostly during the growing season (Caçador et al., 2000; Duarte et al., 2010) and can influence the chemical speciation and mobility of contaminants by changing sediment redox conditions, pH and organic matter content (Castro et al., 2009; Jacob and Otte, 2003; Pedro et al., 2015). Studies on mercury in halophyte colonised salt marsh areas have consistently shown elevated THg and MMHg levels in the rhizosphere, compared to levels found in sediments (Canário et al., 2010), and limited transport of both forms into stems and leaves, (Anjum et al., 2011; Canário et al. 2010; Castro et al., 2009; Válega et al., 2008a, 2008b). These results suggest either reduced translocation to or weak retention of mercury species by aerial parts of

plants, and/or Hg volatilization from these plant organs. In other plant species besides halophytes, most of the Hg content in foliar tissue has been found to come from the atmosphere, being more significant than Hg translocation from the roots (Ericksen et al., 2003; Frescholtz et al., 2003; Gustin et al., 2004; Laacouri et al., 2013; Mao et al., 2013; Marrugo-Negrete et al., 2016a; Tomiyasu et al. 2005). In addition to incorporation into leaves via atmospheric Hg deposition, Hg release from leaves during plant transpiration has also been observed (Weis and Weis, 2004; Windham et al., 2001). Continuous Hg emission from plants into the atmosphere, shown to be positively correlated with air temperature, suggests that Hg translocation within the plants may play a more important role than previously anticipated (Canário et al., 2017). However, the uptake mechanism into the roots and from these organs to the aerial parts of the plants is still not well understood. Although some salt marsh halophytes are able to endure metal/metalloid contamination to a certain degree, excessive levels internalised in the plants can cause severe impairment of fundamental processes linked to protein and energy metabolism (Falchuk et al., 1979; Liu et al., 2013; Patra et al., 2004). In fact, metal/metalloid overload have been shown to trigger severe damage in the photosystem II (PS II) at the biophysical level (Anjum et al., 2016; Duarte et al., 2014; Santos et al., 2014). At the biochemical level, metal/metalloid-accrued increase in reactive oxygen species (ROS) can induce cellular redox homeostasis imbalance (e.g. Anjum et al., 2014b, 2016; Pietrini et al., 2003; Santos et al., 2014, 2015; Schroder et al., 2009). Tolerance mechanisms are developed by the plants in order to escape these harmful effects, for example, metal ions can be prevented from interfering with the cellular metabolism by either cell wall immobilization (Sousa et al., 2008; Wang and Greger, 2004) or chelation with specific peptides or organic acids (Hall, 2002; Yang et al., 2005). Despite these advances, a knowledge gap is noticeable regarding the mechanisms by which THg and MMHg enter and accumulate in plant tissues and how these toxic compounds are transported from roots to the aboveground plant parts. The present work investigates the uptake of mercury species into the roots of salt marsh plants and its subsequent translocation and accumulation by aerial parts (stems and leaves) of the plant, as well as the plants photochemical response to mercury contamination. The study was targeted at the salt marsh plant Halimione portulacoides and aimed at providing critical insight on (i) processes of uptake, translocation and distribution of THg and MMHg within the whole plant, and (ii) toxicity and tolerance mechanisms by investigating plant photochemical responses. In order to achieve these objectives, fully developed halophyte H. portulacoides plants, grown under hydroponic conditions and exposed to mercury stable isotopes, were monitored during a period of 18 days, for changes in (i) T199Hg and MM201Hg content in roots, stems and leaves, (ii) PS II electron transport activity, and (iii) electron transport chain (ETC) behaviour, in leaves. The salt marsh plant H. portulacoides was chosen because it is an abundant halophyte species colonizing salt marshes worldwide, grows well from cuttings, has been shown to accumulate mercury within its root system, and has been suggested as a potential biomonitor for mercury and metal contamination. Hydroponics have already been successfully used to investigate uptake and translocation of mercury species in different environmental compartments of wetland plants (Chattopadhyay et al., 2012; Cui et al., 2014; Rahn, 1973). In the present study, combination of the mercury stable isotope tracer method with hydroponic plant growing was applied, to allow consistent plant exposure to Hg species, thus avoiding


natural environmental mercury bioavailability fluctuations (Greger, 2004). This proved a powerful approach to directly assess short-term (up to 24 h) and long-term (72–432 h) THg and MMHg bioaccumulation and translocation in H. portulacoides. This study contributes to better understand how and to what extent different Hg species (THg and MMHg) enter H. portulacoides and how they are translocated within the plant. 2. Materials and methods 2.1. Sampling area and collection of Halimione portulacoides Sampling of the plant Halimione portulacoides was conducted in a young salt marsh with narrow mudflats, located in the Alcochete area (38°45′38.78″N, 8°56′7.37″W), in the northern part of the Tagus Estuary (Portugal). From the 17.24 km2 of salt marshes existing in Tagus estuary, about 5.32 km2 are colonised by H. portulacoides (Caçador et al., 2013). The young salt marsh is situated within a RAMSAR convention (Ramsar Convention on Wetlands of International Importance especially as Waterfowl Habitat) area, away from urbanized and industrial areas, and was chosen to ensure minimal mercury contamination of the plants. This intertidal salt marsh is inundated twice a day, with the tidal amplitude ranging from about 1 m at neap tide to about 4 m at spring tide. Mercury concentrations in surface sediments of this salt marsh area have been found low, with concentrations of Hg species ranging between 0.20 and 0.60 μg Hg g−1 and 0.20–2.5 ng MMHg g−1 (Canário et al., 2017; Cesário et al., 2016). The H. portulacoides plants were collected, during low tide. Plants of similar size were carefully removed from the sediment, to ensure that the root system was preserved as much as possible, and quickly transported to the laboratory. 2.2. Mercury isotope exposure experiments Once in the laboratory, H. portulacoides plants were carefully washed and let to grow in hydroponic medium containing nutrients for about three weeks, until the root system was fully developed. Fully developed plants were then cultivated in several sets, under hydroponic setting, containing nutrients and 199HgCl2 and MM201HgCl. These isotopes were chosen because they are among the most abundant mercury natural stable isotopes (199Hg: 16.87 (22); 201Hg: 13.18 (9)), and 199Hg has been found the most available natural isotope for methylation whereas MM201Hg degrades similarly to MMHg in natural sediments (Hintelmann et al., 2000). This double spiking approach allowed the simultaneous determination of two processes (methylation and demethylation) in parallel. Experiments were conducted in 50 mL non RackLock DigiTUBEs (SCP Science) made of polypropylene with ultralow catalytic/additive metal content. Each tube contained only one plant individual and 50 mL of spiked growth medium solution. In parallel, plant control sets were grown in hydroponic medium containing only nutrients, under the same experimental setting conditions, for evaluation of photobiological features. The nutrient solution was one quarter strength diluted Hoagland solution with the following composition: 1.25 mM KNO3, 1 mM Ca(NO3)2, 0.5 mM NH4H2PO4, 0.25 mM MgSO4, 50 μM KCl, 25 μMH3BO3, 2 μM MnSO4, 2 μM ZnSO4, 0.5 μM CuSO4, 0.5 μM (NH4)6 Mo7O24 and 20 μM FeNaEDTA. The spiked growth medium solution was prepared by adding 20 μL of a spike solution containing MM201Hg (1.2 μg mL−1) and T199Hg (52.8 μg mL−1) to 1 L of nutrient solution. The concentrations of 199Hg(II) and MM201Hg used were 1056 ng L−1 and 24 ng L−1, respectively, reproducing overlying and pore waters concentrations previously found in a contaminated salt marsh located in the Tagus estuary (Cesário et al., 2016, 2017), nearby this study sampling site. Growth medium solution was changed daily throughout the experiment. Plants were grown inside a FytoScope 130 RGBIR (Photon System Instruments, Czech Republic) chamber programmed with a sinusoidal function simulating a 16 h light:8 h dark cycle provided by a cool white fluorescent light source (maximum

113

light intensity 800, simulating natural light variation from sunrise to sunset). Temperature was set to 20 ± 0.5 °C during day time and 18 ± 0.5 °C at night time. The FytoScope chamber provided air renovation (air flow rate of about 250 L h−1) and circulation as well as even temperature distribution. The plants were sampled at the beginning of the experiment (T0), and after 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days) to allow comparison between short-term (up to 24 h) and long-term (72–432 h) mercury accumulation. Three replicate samples each were obtained for each sampling time. One replicate corresponded to one individual plant. Mercury safety guidelines regarding the handling, storage and waste of mercury solutions were observed throughout the experiments even though elemental Hg was not used, as liquid mercury poses a health hazard because it is volatile and can be absorbed through the skin, reaching the central nervous system, with extremely toxic effects. 2.3. Pulse Amplitude Modulated (PAM) fluorometry analyses Pulse Amplitude Modulated (PAM) chlorophyll fluorescence measurements were performed using a FluoroPen FP100 (Photo System Instruments, Czech Republic), on 30 min dark-adapted control and mercury exposed H. portulacoides plants, as described in Duarte et al. (2014). Effective quantum yields of PS II in the dark reaction centres were determined with a rational saturation pulse method, as described by Schreiber et al. (1986). The measuring light intensity was b0.1 μmol m−2 s−1 of blue light. A 0.8 s saturating blue light pulse with an intensity of 8000 μmol m−2 s−1 was used to estimate the maximum fluorescence signal over time as a function of photon irradiance. The same parameters were measured in light adapted samples, to compare the minimum fluorescence (F′0), the maximum fluorescence (F′M) and the operational photochemical efficiency with dark-adapted samples. The polyphasic rise in fluorescence (OJIP) transient depicts the rate of reduction kinetics of various components of PS II. When dark-adapted cells are illuminated with the saturating light intensity of 3500 μmol m−2 s−1 it exhibits a polyphasic OJIP curve. The distinct inflections in the curve are denoted by the letters in OJIP. The level O represents all the open reaction centres at the onset of illumination with no reduction of QA (fluorescence intensity lasts for 10 ms). The rise of transient from O to J indicates the net photochemical reduction of QA (the stable primary electron acceptor of PS II) to Q− A (lasts for 2 ms). The phase from J to I − − is due to all reduced states of closed RCs such as Q− A QB , QA QB2 and − QA QB H2 (lasts for 2–30 ms). The level P (300 ms) coincides with maximum concentration of Q− A QB2 with plastoquinol pool maximally reduced. The phase P also reflects a balance between light incident at the PS II side and the rate of utilization of the chemical (potential) energy and the rate of heat dissipation (Zhu et al., 2005). Table 1 summarizes all the parameters that were computed from the fluorometric data. 2.4. Sample preparation for mercury isotope analysis Immediately after (PAM) chlorophyll fluorescence measurements, roots, stems and leaves were separated from each plant, thoroughly

Table 1 Fluorometric analysis parameters and their description. Energy fluxes (Kautsky curves)

Description

Area

Corresponds to the oxidized quinone pool size available for reduction and is a function of the area above the Kautsky plot Absorbed energy flux Trapped energy flux Electron transport energy flux Dissipated energy flux Number of available reaction centres per leaf cross section

ABS/CS TR/CS ET/CS DI/CS RC/CS

114


rinsed with distilled water, and maintained at −80 °C. Prior to analyses, the frozen root, stem and leave samples were then dried in an oven at 40 °C for at least 48 h, and reduced to a fine powder in a mortar.

2.5. Total Hg isotope analyses Total Hg (THg) in root, stem and leave samples were measured after digestion of the organic matrix. Samples were weighed into acid washed glass vials, and 100 μL of a 4.5 ng mL−1 solution of enriched 200 Hg2+ were added as internal standard. Samples were then openly digested with 5 mL of an mixture of concentrated nitric acid and sulfuric acid (7:3) in a heating block (120 °C for at least 48 h), with temperature gradually increasing to 160 °C during the last 2 h. After cooling, 100 μL of 2 M BrCl solution were added and digests were diluted to 10 mL with Milli-Q water (18.2 MΩ cm). Immediately before total Hg analysis, 10 μL of a 20% hydroxylamine hydrochloride solution were added to neutralize excess BrCl. Digestion blanks were prepared with every batch of samples. The concentration of Hg isotopes in the samples was quantified using continuous-flow cold-vapor generation with ICP/MS detection (X-Series II, Thermo Fisher). The acidified samples were continuously mixed with a solution of stannous chloride 3% (w/v) in 10% HCl (v/v), the formed mercury vapor was separated from the liquid in a gas-liquid separator (Model L1-2) and the elemental mercury swept into the plasma of the ICP/MS. The following isotopes of Hg were measured: 199Hg (T199Hg spike) and 200Hg (internal standard). Concentrations of individual isotopes were calculated as described in Hintelmann and Ogrinc (2003).

2.6. MMHg isotope analyses For the determination of MMHg, samples were previously distilled to isolate MMHg from the plant matrix. Samples were weighed into Teflon vials and 100 μL of a 104 pg mL−1 solution of enriched MM202Hg were added to the samples as an internal standard to correct for procedural losses. Subsequently, 10 mL of Milli-Q water (18.2 MΩ cm), 500 μL of 9 M H2SO4 and 200 μL of a 20% KCl solution were added to the sample Teflon vials. Receiving vials were filled with 5 mL of Milli-Q water and vials were closed and connected with Teflon tubes. The distillation vials were placed into a heating block (140 °C), and MMHg was distilled from the samples under a supporting nitrogen stream (50 mL min−1) to the receiving vials. Distillation was stopped when approximately 85% of the original volume was transferred into the receiving vials. Concentration of MMHg in distillates was determined by species-specific isotope dilution inductively coupled plasma mass spectrometry according to Hintelmann and Evans (1997) and Hintelmann and Ogrinc (2003) using an automated Tekran 2700 system coupled to ICP/MS (X-Series II, Thermo-Fisher). The following isotopes of Hg were measured: MM201Hg (MM201Hg spike) and MM202Hg (internal standard). Peak areas were used for quantification, and concentrations of individual isotopes were calculated according to Hintelmann and Ogrinc (2003).

2.8. Statistical analysis Due to the lack of normality and homogeneity, the data statistical analysis was based on non-parametric tests. In order to compare the differences between mercury concentrations in roots, stems and leaves, a Kruskal-Wallis analysis of variance was performed using Statistica Software (Statsoft). Significance was assumed when p b 0.05. Principal Component Analysis (PCA) was applied to a matrix of 4 variables, namely Photosynthetically Active Radiation (PAR) and temperature (measured in the growth chamber), and THg and MMHg plant content, and 27 objects (root, stem and leaf samples collected at 0, 1, 2, 4, 24, 72, 120, 168 and 432 h of mercury exposure). The software used was NTSYS-PC (Numerical Taxonomy and Multivariate System Analysis) Version 2.0 software package. 3. Results 3.1. Distribution of THg in the H. portulacoides plant organs Time variation of the concentrations of 199Hg in the roots, stems and leaves of H. portulacoides plants is shown in Fig. 1. In roots, concentration of 199Hg increased steadily from 2.7 ± 0.21 to 3711 ± 410 pg g−1 (d.w.) throughout the entire exposure period. In stems and leaves, an increase in concentration of 199Hg was found until day 7 (168 h), followed by a slight decrease at day 18 (432 h). In stems, values increased from 0.12 ± 0.022 to 62 ± 5.2 pg g−1 (d.w.), then lowering to 56 ± 10.1 pg g−1 (d.w.). In leaves, 199Hg levels raised from 0.49± 0.020 to 25 ± 8.2 pg g−1 (d.w.) and then reduced to 16 ± 10.4 pg g−1 (d.w.). Concentrations of 199Hg were significantly higher (p b 0.05) in roots than in stems and leaves, for each sampling time. Average 199Hg value in roots was 1076 pg g−1 (d.w.) whereas stems and leaves values were 21 and 8 pg g−1 (d.w.), respectively. A significantly positive correlation regarding 199Hg concentrations was found between roots and stems (r = 0.93, p b 0.01, n = 9), and between stems and leaves (r = 0.94, p b 0.01, n = 9). 3.2. Distribution of MMHg in the H. portulacoides plant organs Time variation of the concentrations of MM201Hg in the roots, stems and leaves of H. portulacoides plants is shown in Fig. 2. During the exposure period, isotope concentrations in the roots significantly increased (p b 0.05) from 0.61 ± 0.20 pg g−1 (d.w.), at the beginning of the experiment, to 807 ± 350 pg g−1 (d.w.) at day 5 (120 h), significantly decreasing (p b 0.05) to 388 ± 50 pg g−1 (d.w.) at day 18 (432 h). In stems and leaves, the MM201Hg temporal variation was somewhat different, values raised until day 7 (168 h), showing only a slight nonsignificant decrease at the end of the exposure period. The highest levels found were 12 ± 2.8 and 5.8 ± 0.7 μg g−1 (d.w.) for stems and leaves, respectively. Concentrations of MM201Hg varied within the plants, with roots presenting significantly higher (p b 0.05) levels than stems and leaves (Fig. 2). Average MM201Hg concentration in roots was 288 pg g−1 (d.w.), in contrast with 4.4 and 2.1 pg g−1 (d.w.) found in stems and leaves, respectively. The concentrations of MM201Hg were significantly correlated between roots and stems (r = 0.87, p b 0.01, n = 9), and between stems and leaves (r = 0.99, p b 0.01, n = 9).

2.7. Accumulation and translocation factors The root accumulation factor (AF) in H. portulacoides plants was calculated as the ratio between mercury (THg or MMHg) concentration in roots and its concentration in the growth medium, according to (Matache et al., 2013). Translocation factors (TF) of THg and MMHg, between the different plant organs, were calculated by the ratio of [metalloid]stems/[metalloid] roots (TF S/R), [metalloid]leaves/[metalloid]stems (TF L/S), and [metalloid] leaves/[metalloid]roots (TF L/R), expressing the metalloid translocation between the plant organs (Deng et al., 2004).

3.3. Effect of THg, MMHg, temperature and radiation on mercury bioaccumulation in the H. portulacoides plant organs A PCA was performed to highlight possible relationships between roots, stems and leaves over the exposure period, and THg, MMHg, temperature and radiation (PAR), and thus evaluate the potential of Hg release by volatilization through leaves associated with temperature and PAR (Fig. 3). The PC1 explained 56% of the variance and separated roots from stems and leaves, due to a combination of 199Hg and MM201Hg content, temperature and radiation. Roots gathered in one


115

4500 4000

(pg g-1)

g

400

3500

199Hg

H. portulacoides roots

Short-term

600

200

3000

f

0 0

6

2500

12 18 Time (hours)

24

f

2000 1500

e

1000

e

500 d 0 70

bc a

0

50

10

150 h

100

60

(pg g-1) 199Hg

300

350

400

h

H. portulacoides stems

50 0 0

40

6

12 18 Time (hours)

24

g

30 20

f

10 d

e bc

0 a 70 0 3 60

(pg g-1)

250

Time (hours)

5

199Hg

200

50

100

150

200

250

Time (hours)

2

300

350

400

H. portulacoides leaves

1

50

0 0

40

6

12 18 Time (hours)

24

c

30

c

c

20 10 0

bb b a

0

b

a 50

100

150

200

250

300

350

400

450

Time (hours) Fig. 1. Concentrations of T199Hg (pg g−1 d.w.) in Halimione portulacoides roots, stems and leaves, at the beginning of the exposure experiment (0 h), and after 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days) of exposure to both THg and MMHg combined (mean ± standard deviation, n = 3 replicate roots, stems or leaves from one plant).

group (except for roots at T0 with low mercury content similar to that found in stems and roots) were projected in the opposite side to stems and leaves, highlighting roots as the main organs for mercury accumulation. The PCA separated stem and leaf samples collected during the first 24 h of mercury exposure (short-term uptake) from those obtained after a longer period of exposure (long-term uptake), revealing that the 199Hg and MM201Hg long-term uptake in stems and leaves is more effected by temperature and PAR than short-term uptake. Stems and leaves were projected on the opposite side to both mercury forms, and close to temperature and PAR, on the PC2 axis (43%), which emphasizes that mercury content in the aerial parts was more influenced by temperature and radiation, apart from the lower mercury content. 3.4. Bioaccumulation and translocation of mercury by H. portulacoides Accumulation factors (roots/growth medium) and translocation factors of THg and MMHg between the different plant organs (translocation factors: stems/roots; leaves/roots; leaves/stems) in H. portulacoides plants are shown in Table 2. The bioaccumulation factors for THg and MMHg were always higher than one (1.02 ± 1.2 for THg, 12 ± 11 for MMHg), except at the beginning of the experiment, which highlights H. portulacoides as an accumulator of mercury in

roots. The translocation factors obtained were always lower than one for both THg and MMHg, indicating that this species does not accumulate high amounts of either form of mercury in the aerial organs. Significant differences in translocation were noticed between the different plant organs. Average translocation from roots to stems (0.022 ± 0.011 for THg, 0.12 ± 031 for MMHg) and from roots to leaves (0.027 ± 0.059 for THg, 0.042 ± 0.11 for MMHg) were significantly lower (p b 0.05) than that found from stems to leaves (0.78 ± 0.14 for THg, 0.36 ± 0.19 for MMHg). 3.5. Pulse Amplitude Modulated (PAM) fluorometry The PAM fluorometry highlighted changes in the energy transduction fluxes, produced by exposure to both THg and MMHg combined on the H. portulacoides plants, although a clear change pattern was not observed over time (Fig. 4). An increase in the number of reaction centres (RC) available for reduction per cell cross-section was found. This feature allowed a higher amount of effectively absorbed photon-flux by PS II antenna chlorophyll (ABS/CS). Changes on the maximal energy flux which entered the RC of PS II and reduced QA to Q− A (TR/CS) were also observed. An enhancing effect on the maximal electron transport (ET/CS) from Q− A to the PS I and consequent re-reduction the photooxidized RC of PS I, was visible mostly at the beginning of exposure (1

116


1400 300

Short-term

MM201Hg (pg g-1)

1200

H. portulacoides roots

e

200

100

1000

0 0

800

6

12 18 Time (hours)

e

24

600 d d

400 d c

200

MM201Hg (pg g-1)

0 16 0

c b a

50

3

14

2

12

1

10

0

100

150

c

200

250

Time (hours)

300

350

400

H. portulacoides stems

c

c 0

6

12 18 Time (hours)

24

8 6 b

4 b

2 0 9 0 8

b b a

50

2

100

150

200

1

7

MM201Hg (pg g-1)

b

250

300

350

400

Time (hours)

H. portulacoides leaves c

200

300

c

6

0 0

6

5

12 18 Time (hours)

24

c

4 3 2

a b a a a

1 0

0

b

50

100

150

250

350

400

450

Time (hours) Fig. 2. Concentrations of MM201Hg (pg g−1 d.w.) in Halimione portulacoides roots, stems and leaves, at the beginning of the exposure experiment (0 h), and after 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days) of exposure to both THg and MMHg combined (mean ± standard deviation, n = 3 replicate roots, stems or leaves from one plant).

1.0

PC2 (43%)

Roots Stems Leaves

MMHg THg

and 4 h), contrasting with decreases at 72 and 432 h. Concomitantly, an increase on the dissipated energy flux (DI/CS) was also noticed. Leaves showed a significant increase in the size of the quinone pool available for reduction, compared to controls, after 120 h of exposure (Fig. 5).

0.5

-1.0

-0.5

0.0

0.0

0.5

1.0

PC1 (56%)

1.5

T72-T432 -0.5

PAR Temp

T0-T24 -1.0

Fig. 3. Projection of Photosynthetically Active Radiation (PAR) and temperature (both measured in the growth chamber), and THg and MMHg plant content, and 27 samples (roots, stems and leaves) collected at 0, 1, 2, 4, 24, 72, 120, 168 and 432 h of exposure to both 199THg and MM201Hg combined, obtained from the Principal Component Analysis (PCA). Short-term (up to 24 h) and long-term (72–432 h) stem and leaf samples are highlighted in oval shapes. Percentage of total variance is indicated in brackets close to principal component axes.

Table 2 Ranges and average of bioaccumulation factor (roots/growth medium) and translocation factors (translocation factors: stems/roots; leaves/roots; leaves/stems), obtained during the exposure experiment, between the different plant organs of THg and MMHg, within H. portulacoides plants exposed to 199Hg(II) (1056 ng L−1) and MM201Hg (24 ng L−1) combined (n = 3; average ± SD). Metal

199

Bioaccumulation factor in roots Translocation factor stems/roots (TF S/R) Translocation factor leaves/roots (TF L/R) Translocation factor leaves/stems (TF L/S)

0.0025 ± 0.00020–3.5 ± 0.39 1.02 ± 1.2 0.013 ± 0.00010–0.028 ± 0.0052 0.022 ± 0.011 0.17 ± 0.11–4.03 ± 0.56 0.027 ± 0.059 0.0023 ± 0.0018–0.18 ± 0.0070 0.78 ± 0.14

Hg(II)

MM201Hg 0.025 ± 0.0083–34 ± 15 12 ± 11 0.0072 ± 0.0018–0.99 ± 0.17 0.12 ± 031 0.038 ± 0.0055–0.59 ± 0.025 0.042 ± 0.11 0.00027 ± 0.000030–0.34 ± 0.030 0.36 ± 0.19


117

100

ABS/CS

RC/CS

TR/CS

ET/CS

DI/CS

RC/ABS

Medium deviations (%) from control average of phenomological variable s

50

0

-50

-100 100

50

0

-50

-100 100

50

0

-50

-100 1

2

4

24

72

120

168

432

1

2

4

24

72

120

168

432

Exposure time (hours) Fig. 4. Medium deviations (%) of 30 min dark-adapted exposed leaves, from the average of 30 min dark-adapted control leaves, of phenomological variables of Halimione portulacoides, at 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days) of exposure to both 199THg and MM201Hg combined (mean ± standard deviation, n = 3).

4. Discussion

Medium deviations (%) from control average of quinone pool size

The mercury stable isotope tracer approach combined with hydroponic growing of the salt marsh plant Halimione portulacoides exposed to THg and MMHg provides a powerful method to directly assess

100

Quinone Pool Size

50 0 -50 -100 1

2

4

24

72

120

168

432

Exposure time (hours)

Fig. 5. Medium deviations (%) of 30 min dark-adapted exposed leaves, from the average of 30 min dark-adapted control leaves, of quinone pool size of Halimione portulacoides at 1, 2, 4, 24, 72, 120, 168 (7 days) and 432 h (18 days) of exposure to both 199THg and MM201Hg combined (mean ± standard deviation, n = 3).

bioaccumulation and translocation mechanisms in the plant. This method provided a consistent exposure to Hg species over time, overcoming challenges often associated with changes in the bioavailability of mercury to salt marsh plants in the natural environment (Greger, 2004), and allowed the determination of the actual levels of both THg and MMHg accumulation and translocation in the plants. The use of hydroponic growing allows Hg to be readily available for uptake under optimal controlled and homogeneous nutrient conditions for plant growth (Carrasco-Gil et al., 2013; Chattopadhyay et al., 2012), thus reducing variability in the responses of plants to Hg uptake, and also enables the detection of low isotope concentrations. The concentrations of 199Hg(II) and MM201Hg found within the plant organs (roots, stems and leaves) clearly showed that both forms of mercury were not only available for root uptake, but were also transported to the aerial parts (stems and leaves) of the plant. The hydroponic growth system has been found to facilitate translocation to leaves which might be associated with root architecture with fewer apoplastic barriers (i.e. Casparian bands and suberin lamellae), compared to plants grown in contaminated soils displaying more apoplastic barriers (Redjala et al., 2011). This way it was possible to clarify if translocation to leaves was a valid pathway in H. portulacoides. Levels of T199Hg and MM201Hg in roots significantly exceeded the concentrations found in stems and leaves. These results corroborate previous studies showing that roots are the main sites of mercury retention for H. portulacoides collected directly from salt marshes (Anjum et al., 2011; Canário et al. 2007, 2010; Castro et al., 2009; Válega et al. 2008a, b), other salt

118


marshes species grown under hydroponic conditions in the presence of HgCl2 (Chattopadhyay et al., 2012; Cui et al., 2014; Rahn, 1973), as well as for other species (Marrugo-Negrete et al., 2016b). In general, roots from salt marsh species have been shown to accumulate comparatively much more metals than any other aerial organ (Almeida et al., 2006; Peverly et al., 1995; Sousa et al., 2008). The higher levels of both THg and MMHg found in the roots in comparison with stems and leaves appeared to reflect the role of the plant organs and the allocation strategy used by plants to cope with mercury stress. Roots are storage organs which can take up mercury from the surrounding environment via the same uptake processes used for essential micronutrient metal ions (Patra et al., 2004). In addition, there is high affinity of both THg and MMHg cations for sulphydryl (–SH) groups. In fact, effective uptake of THg and MMHg by the roots has been shown to occur by sequestration into two classes of cysteine-rich peptides, metallothioneins and phytochelatins (Clarkson, 1993; Skinner et al., 2007), through binding to organic sulphur (R-SH) groups on the cysteine residues in those peptides being glutathionated and transported into vacuoles for long term sequestration (Brouwer et al., 1993; Patra et al., 2004). Alternatively, they can be transported into the aerial parts of the plant following osmotic passage into xylem (Weis and Weis, 2004), which seemingly was the case to a limited extent in H. portulacoides, as the levels of both mercury forms found in stems and leaves were relatively low. This may be partially due to the fact that stems and leaves are not storage organs. Instead, they are a transport organ for fluids (stems) and key metabolic sites mostly dedicated to photosynthesis (leaves). It has been shown that metals are preferably stored in vacuoles and in the cell wall, to preserve metabolic active areas (such as chloroplast and mitochondria) reducing the metal toxicity in the plant (Küpper et al., 2001; Psaras et al., 2000). Below-ground plant cells generally lack chloroplasts, which are prevalent in leaves and in the cells near the surface of green stems, which may have contributed to the low levels of mercury measured in these organs. Furthermore, translocation of both THg and MMHg was found mainly between growth medium and roots with marginal amounts translocated to the aboveground organs. Previous studies focusing on H. portulacoides have shown similar translocation factor values (Castro et al., 2009; Sousa et al., 2008), suggesting limited mobility of the metals once inside the plant and corroborating the fact that mercury is predominantly accumulated in the roots rather than in the above-ground part of this salt marsh plant (Canário et al., 2010; Castro et al., 2009; Sousa et al., 2008; Válega et al., 2008a,b). However, the presence of both mercury isotopes in the stems and leaves and the highly significant correlations found between roots and stems, and stems and leaves, for both THg and MMHg concentrations, indicate that mercury concentrations in the leaves were due to translocation from the roots and show that direct mercury translocation between the below- and above-ground plant is a valid pathway. Therefore, the possibility of mercury being released from the leaves themselves due to evapotranspiration cannot be excluded, and maybe another factor explaining the large difference of mercury content between below- and above-ground tissues. Results from the PCA (Fig. 3) showing that mercury content in the aerial parts of H. portulacoides was influenced by temperature and radiation supported the possibility of mercury release by stems and leaves (Canário et al., 2017; Sizmur et al. 2017; Weis and Weis, 2004; Windham et al., 2001). Although correlation between Hg fluxes and temperature may eventually reflect the correlation between mercury flux and solar radiation, and mercury volatilization has been found independent of air temperature in a previous study (Smith and Reinfelder, 2009), positive relationships between Hg flux and both solar radiation and temperature were observed in plant colonised salt marsh areas (Sizmur et al. 2017) as well as in subtropical forested areas (Ma et al., 2013). Nevertheless, correlation between Hg flux and radiation indicated that emission of Hg(0) is due to photoreduction of Hg(II) and ensuing volatilization (Lee et al., 2000; Smith and Reinfelder, 2009). Mercury volatilization may have been related to stomatal aperture and subsequent enhancement in plant transpiration (Pezeshki, 2001;

Pezeshki and DeLaune, 2012) under higher temperature and radiation, as increases in these two environmental parameters are known to induce stomatal opening (e.g. Araújo et al., 2011; Shimazaki et al., 2007; Urban et al., 2017). The PCA also highlighted that the 199Hg and MM201Hg long-term uptake in stems and leaves, leading to higher mercury content, was more influenced by temperature and radiation than short-term uptake. The fact that leaf Hg concentration has been found highly positively correlated with Hg release (Windham et al., 2001), in association with the effect of temperature and radiation on stomatal aperture (Araújo et al., 2011; Shimazaki et al., 2007; Urban et al., 2017), further supports that both 199Hg and MM201Hg were being released by stems and leaves during the exposure period, and possibly more intensely during long-term exposure. This is also reinforced by Canário et al. (2017) findings, showing a vegetation to air elemental mercury (Hg(0)) flux of 0.48 ± 0.40 ng Hg m−2 h−1 for H. portulacoides, which indicates the existence of leave release mechanisms enabling the plants to eliminate toxic THg and MMHg via stomatal movements. This was further supported by the observed diurnal pattern of elevated daylight and lower nighttime fluxes (Canário et al. 2017), as stomatal aperture is generally increased by light and declines in the dark (Shimazaki et al., 2007). Furthermore, Canário et al. (2017) also observed the absence of negative fluxes (air to vegetation) in this salt marsh plant, which suggests that deposition of mercury that might have accumulated within the growth chamber, with consequent absorption by the leaves, did not occur and did not play a role in mercury accumulation by this species for this experimental setting. Although in other plant species besides halophytes, most of the Hg input to leaves has been found to come from the atmosphere whereas root levels reflect soil or growth medium Hg concentrations (Ericksen et al., 2003; Frescholtz et al., 2003; Gustin et al., 2014; Laacouri et al., 2013; Mao et al., 2013; Marrugo-Negrete et al., 2016a; Tomiyasu et al. 2005), the aeration and circulation provided by the growth chamber used in this study may have prevented a Hg flux from air to leaves. The dose of THg and MMHg applied for the experiments with H. portulacoides, did not lead to typical toxicity symptoms during the first hours of exposure, as seen by an efficient ET/CS energy flux and low energy dissipation. However, after 120 h of exposure, some signs of stress were evident. The size of the oxidized quinone pool showed an increase, indicative of low electron transport rate from PS II to PS I, also observable by a low ET/CS. This is a typical sign of stress under metal/metalloid exposure due to the redox interference of the cations inside the chloroplasts and the oxidative burst generated by the metal induced ROS and consequent membrane lipid peroxidation, where the quinones and the whole ETC are anchored (Anjum et al., 2016). Inevitably, this leads to an increase in the energy of dissipation, making the chloroplast less efficient, and thus producing lower amounts of energy to the whole plant (Anjum et al., 2016; Santos et al., 2014, 2015). Although there is a good translocation rate from the roots and stems to the leaves, this may come at a cost to the plant. Possible mercury volatilization through stomata implicates an efficient mass flow transport from roots to leaves, which have additional energetic costs due to the partial impairment of the ETC and consequent lower conversions of light energy into chemical energy to supply the needs of the plant. Nevertheless, membrane and biochemical adaptations are often exhibited by plants growing in the field that allow them to cope with chronic metal exposure (Duarte et al., 2013). 5. Conclusions Overall, these results indicate that mercury is mainly accumulated into the roots of H. portulacoides, and direct translocation to the aerial parts of the plant actually occurs. Evapotranspiration of mercury from the leaves of the plant possibly explains differences between mercury content between below- and above-ground tissues, and further work is needed to confirm and evaluate the magnitude of this process. Even though at low levels, mercury can impact photochemistry with


prolonged exposure, evidencing the toxicity of both metalloid forms, THg and MMHg to this salt marsh plant. Acknowledgements This work was performed under the project PLANTA — Effect of saltmarsh plants on mercury methylation, transport and volatilization to the atmosphere (PTDC/AAC-AMB/115798/2009), funded by the Portuguese Foundation for Science and Technology (FCT). M.T. Cabrita, B. Duarte and R. Cesário wish to acknowledge grants by FCT (Grants SFRH/BPD/50348/2009 and SFRH/BPD/115162/2016, and SFRH/BD/ 86441/2012, respectively). The authors would like to thank the anonymous reviewers for their constructive comments, which helped improving the manuscript. References Almeida, C.M.R., Mucha, A.P., Vasconcelos, M.T.S.D., 2006. Comparison of the role of the sea club-rush S. maritimus and the sea rush J. maritimus in terms of concentration, speciation and bioaccumulation of metals in the estuarine sediment. Environ. Pollut. 142, 151–159. Anjum, N.A., Ahmad, I., Válega, M., Pacheco, M., Figueira, E., Duarte, A.C., Pereira, E., 2011. Impact of seasonal fluctuations on the sediment-mercury, its accumulation and partitioning in Halimione portulacoides and Juncus maritimus collected from Ria de Aveiro coastal lagoon (Portugal). Water Air Soil Pollut. 222, 1–15. Anjum, N.A., Ahmad, I., Valega, M., Mohmood, I., Gill, S.S., Tuteja, N., Armando, C., Duarte, A.C., Pereira, E., 2014a. Salt marsh halophyte services to metal-metalloid remediation: assessment of the processes and underlying mechanisms. Crit. Rev. Environ. Sci. Technol. 44, 2038–2106. Anjum, N.A., Israr, M., Duarte, A.C., Pereira, M.E., Ahmad, I., 2014b. Halimione portulacoides (L.) physiological/biochemical characterization for its adaptive responses to environmental mercury exposure. Environ. Res. 131, 39–49. Anjum, N.A., Duarte, B., Caçador, I., Sleimi, N., Duarte, A.C., Pereira, E., 2016. Biophysical and biochemical markers of metal/metalloid-impacts in salt marsh halophytes and their implications. Front. Environ. Sci. 4, 24. Araújo, W.L., Fernie, A.R., Nunes-Nesi, A., 2011. Control of stomatal aperture. A renaissance of the old guard. Plant Signal. Behav. 6, 1305–1311. Barbier, E.B., Hacker, S.D., Kennedy, C., Koch, E.W., Stier, A.C., Silliman, B.R., 2011. The value of estuarine and coastal ecosystem services. Ecol. Monogr. 81, 169–193. Brouwer, M., Hoexum-Brouwer, T., Cashon, R.E., 1993. A putative glutatione-binding site in Cd, Zn-metallothionein identified by equilibrium binding and molecular-modelling studies. Biochem. J. 294, 219–225. Burgess, N.M., Meyer, M.W., 2008. Methylmercury exposure associated with reduced productivity in common loons. Ecotoxicology 17, 83–91. Caçador, I., Vale, C., Catarino, F., 1993. Effects of plants on the accumulation of Zn, Pb, Cu and Cd in sediments of the Tagus estuary salt marshes, Portugal. Stud. Environ. Sci. 55, 355–364. Caçador, I., Vale, C., Catarino, F., 1996. The influence of plants on concentration and fractionation of Zn, Pb, and Cu in salt marsh sediments (Tagus Estuary, Portugal). J. Aquat. Ecosyst. Health 5, 193–198. Caçador, I., Vale, C., Catarino, F., 2000. Seasonal variation of Zn, Pb, Cu and Cd concentrations in the root-sediment system of Spartina maritima and Halimione portulacoides from Tagus estuary salt marshes. Mar. Environ. Res. 49, 279–290. Caçador, I., Caetano, M., Duarte, B., Vale, C., 2009. Stock and losses of trace metals from salt marsh plants. Mar. Environ. Res. 67 (2), 75–82. Caçador, I., Neto, J.M., Duarte, B., Barroso, D.V., Pinto, M., Marques, J.C., 2013. Development of an Angiosperm Quality Assessment Index (AQuA-Index) for ecological quality evaluation of Portuguese water bodies — a multi-metric approach. Ecol Indic. 25, 141–148. Canário, J., Vale, C., Caetano, M., 2005. Distribution of monomethylmercury and mercury in surface sediments of the Tagus Estuary (Portugal). Marine Poll Bull 50, 1121–1145. Canário, J., Caetano, M., Vale, C., Cesario, R., 2007. Evidence for elevated production of methylmercury in salt marshes. Environ. Sci. Technol. 41, 7376–7382. Canário, J., Vale, C., Poissant, L., Nogueira, M., Pilote, M., Branco, V., 2010. Mercury in sediments and vegetation in a moderate contaminated salt-marsh (Tagus Estuary, Portugal). J. Environ. Sci. 22 (8), 1151–1157. Canário, J., Poissant, L., Pilote, M., Caetano, M., Hintelmann, H., O'Driscoll, N.J., 2017. Saltmarsh plants as potential sources of Hg0 into the atmosphere. Atmos. Environ. 152, 458–464. Carrasco-Gil, S., Siebner, H., Leduc, D.L., Webb, S.M., Millán, R., Andrews, J.C., Hernández, L.E., 2013. Mercury localization and speciation in plants grown hydroponically or in a natural environment. Environ. Sci. Technol. 47 (7), 3082–3090. Castro, R., Pereira, S., Lima, A., Corticeiro, S., Válega, M., Pereira, E., Duarte, A., Figueira, E., 2009. Accumulation, distribution and cellular partitioning of mercury in several halophytes of a contaminated salt marsh. Chemosphere 76 (10), 1348–1355. Cesário, R., Monteiro, C.E., Nogueira, M., O'Driscoll, N.J., Caetano, M., Hintelmann, H., Mota, A.M., Canário, J., 2016. Mercury and methylmercury dynamics in sediments on a protected area of Tagus estuary (Portugal). Water Air Soil Pollut. 227, 475. Cesário, R., Poissant, L., Pilote, M., O'Driscoll, N.J., Mota, A.M., Canário, J., 2017. Dissolved gaseous mercury formation and mercury volatilization in intertidal sediments. Sci. Total Environ. 603–604, 279–289.

119

Chattopadhyay, S., Fimmen, R.L., Yates, B.J., Lal, V., Randall, P., 2012. Phytoremediation of mercury- and methyl mercury-contaminated sediments by water hyacinth (Eichhornia crassipes). Int. J. Phytorem. 14, 142–161. Clarkson, T.W., 1993. Molecular and ionic mimicry of toxic metals. Annu. Rev. Pharmacol. Toxicol. 33, 545–571. Clayden, M.G., Kidd, K.A., Wyn, B., Kirk, J.L., Muir, D.C.G., O'Driscoll, N.J., 2013. Mercury biomagnification through food webs is affected by physical and chemical characteristics of lakes. Environ. Sci. Technol. 47 (21), 12047–12053. Costanza, R., d'Arge, R., de Groot, R., Farberk, S., Grasso, M., Hannon, B., Limburg, K., Naeem, S., O'Neill, R.V., Paruelo, J., Raskin, R.G., Suttonkk, P., van den Belt, M., 1997. The value of the world's ecosystem services and natural capital. Nature 387, 253–260. Cui, L., Feng, X., Lin, C.-J., Wang, X., Meng, B., Wang, X., Wang, H., 2014. Accumulation and translocation of 198Hg in four crop species. Environ. Toxicol. Chem. 33 (2), 334–340. Deng, H., Ye, Z.H., Wong, M.H., 2004. Accumulation of lead, zinc, copper and cadmium by 12 wetland plant species thriving in metal-contaminated sites in China. Environ. Pollut. 132 (1), 29–40. Driscoll, C.T., Mason, R.P., Chan, H.M., Jacob, D.J., Pirrone, N., 2013. Mercury as a global pollutant: sources, pathways, and effects. Environ. Sci. Technol. 47 (10), 4967–4983. Duarte, B., Caetano, M., Almeida, P., Vale, C., Caçador, I., 2010. Accumulation and biological cycling of heavy metal in the root-sediment system of four salt marsh species, from Tagus estuary (Portugal). Environ. Pollut. 158, 1661–1668. Duarte, C., Losada, I.J., Hendriks, I.E., Mazarrasa, I., Marbà, N., 2013. The role of coastal plant communities for climate change mitigation and adaptation. Nat. Clim. Change 3, 961–968. Duarte, B., Santos, D., Marques, J.C., Caçador, I., 2014. Biophysical probing of Spartina maritima photo-system II changes during increased submersion periods: possible adaptation to sea level rise. Plant Physiol. Biochem. 77, 122–132. Ericksen, J.A., Gustin, M.S., Schorran, D.E., Johnson, D.W., Lindberg, S.E., Coleman, J.S., 2003. Accumulation of atmospheric mercury in forest foliage. Atmos. Environ. 37 (12), 1613–1622. Evers, D.C., Han, Y.-J., Driscoll, C.T., Kamman, N.C., Goodale, W.M., Lambert, K.F., Holsen, T.M., Chen, C.Y., Clair, T.A., Butler, T.J., 2007. Biological mercury hotspots in the Northeastern United States and Southeastern Canada. Bioscience 57 (1), 1–15. Falchuk, K.H., Goldwater, L.J., Vallee, B.L., 1979. The biochemistry and toxicology of mercury. Macmillan, New York, pp. 261–284. Fitzgerald, W.F., Lamborg, C.H., 2003. Geochemistry of mercury in the environment. Treatise Geochem. 9, 107–148. Frescholtz, T.E., Gustin, M.S., Schorran, D.E., Fernandez, G.C., 2003. Assessing the source of mercury in foliar tissue of quaking aspen. Environ. Toxicol. Chem. 22, 2114–2119. Giblin, A.E., Weider, R.K., 1992. Sulfur cycling in marine and freshwater wetlands. In: Howarth, R.W., Stewart, J.W.B., Ivanov, M.V. (Eds.), SCOPE 48 — Sulphur Cycling on the Continents, Chapter 5. SCOPE. Wiley, U.K. Greenberg, R., Maldonado, J.E., Droege, S., McDonald, M.V., 2006. Tidal marshes: a global perspective on the evolution and conservation of their terrestrial vertebrates. BioScience 56 (8), 675–685. Greger, M., 2004. Metal availability, uptake, transport and accumulation in plants. In: Prasad, M.N.V. (Ed.), Heavy Metal Stress in Plants — From Biomolecules to Ecosystems, second ed. Springer-Verlag, Heidelberg, Berlin, Germany, pp. 1–27. Gupta, M., Chandra, P., 1998. Bioaccumulation and toxicity of mercury in rootedsubmerged macrophyte Vallisneria spiralis. Environ. Pollut. 103, 327–332. Gustin, M.S., Ericksen, J.A., Schorran, D.E., Johnson, D.W., Lindberg, S.E., Coleman, J.S., 2014. Application of controlled mesocosm for understanding mercury plant-soil-air exchange. Environ. Sci. Technol. 38, 6044–6050. Gworek, B., Bemowska-Kałabun, O., Kijeńska, M., Wrzosek-Jakubowska, J., 2016. Mercury in marine and oceanic waters—a review. Water Air Soil Pollut. 227, 371. Hall, J.L., 2002. Cellular mechanisms for heavy metal detoxification and tolerance. J. Exp. Bot. 53, 1–11. Hintelmann, H., Evans, R.D., 1997. Application of stable isotopes in environmental tracer studies — measurement of monomethylmercury (CH3Hg+) by isotope dilution ICP-MS and detection of species transformation. Fresenius J. Anal. Chem. 358 (3), 378–385. Hintelmann, H., Ogrinc, N., 2003. Determination of stable mercury isotopes by ICP/MS and their application in environmental studies. Biogeochemistry of Environmentally Important Trace Elements. ACS Symposium Series 835, pp. 321–338. Hintelmann, H., Keppel-Jones, K., Evans, R.D., 2000. Constants of mercury methylation and demethylation rates in sediments and comparison of tracer and ambient mercury availability. Environ. Toxicol. Chem. 19, 2204–2211. Hong, Y.-S., Kim, Y.-M., Lee, K.-E., 2012. Methylmercury exposure and health effects. J. Prev. Med. Public Health 45 (6), 353–363. Hung, G.A., Chmura, G.L., 2006. Mercury accumulation in surface sediments of salt marshes of the Bay of Fundy. Environ. Pollut. 142, 418–431. Jacob, D.L., Otte, M.L., 2003. Conflicting processes in the wetland plant rhizosphere: metal retention or mobilization? Water Air Soil Pollut. 3, 91–104. Küpper, H., Lombi, E., Zhao, F., Wieshammer, G., McGrath, S.P., 2001. Cellular compartmentation of nickel in the hyperaccumulators Alyssum lesbiacum, Alyssum bertolonii and Thlaspi goesingense. J. Exp. Bot. 52, 2291–2300. Laacouri, A., Nater, E.A., Kolka, R.K., 2013. Distribution and uptake dynamics of mercury in leaves of common deciduous tree species in Minnesota, U.S.A. Environ. Sci. Technol. 47, 10462–10470. Lee, X., Benoit, G., Hu, X., 2000. Total gaseous mercury concentration and flux over a coastal saltmarsh vegetation in Connecticut, USA. Atmos. Environ. 34, 4205–4213. Lindqvist, O., Johansson, K., Bringmark, L., Timm, B., Aastrup, M., Andersson, A., Hovsenius, G., Hakanson, L., Iverfeldt, A., Meili, M., 1991. Mercury in Swedish environments. Water Air Soil Pollut. 55, 23–30. Liu, X., Wu, H., Ji, C., Wei, L., Zhao, J., Yu, J., 2013. An integrated proteomic and metabolomic study on the chronic effects of mercury in Suaeda salsa under an environmentally relevant salinity. PLoS One 8 (5), e64041.

120


Ma, M., Wang, D., Sun, R., Shen, Y., Huang, L., 2013. Gaseous mercury emissions from subtropical forested and open field soils in a national nature reserve, southwest China. Atmos. Environ. 346 (64), 116–123. Mao, Y., Li, Y., Richards, J., Cai, Y., 2013. Investigating uptake and translocation of mercury species by sawgrass (Cladium jamaicense) using a stable isotope tracer technique. Environ. Sci. Technol. 47 (17), 9678–9684. Marrugo-Negrete, J., Marrugo-Madrida, S., Pinedo-Hernández, J., Durango-Hernández, J., Díez, S., 2016a. Screening of native plant species for phytoremediation potential at a Hg-contaminated mining site. Sci. Total Environ. 542 (Part A), 809–816. Marrugo-Negrete, J., Durango-Hernández, J., Pinedo-Hernández, J., Enamorado-Montes, G., Díez, S., 2016b. Mercury uptake and effects on growth in Jatropha curcas. J. Environ. Sci. 48, 120–125. Matache, M.L., Marin, C., Rozylowicz, L., Tudorache, A., 2013. Plants accumulating heavy metals in the Danube River wetlands. J. Environ. Health Sci. Eng. 11, 39. Mcowen, C., Weatherdon, L., Bochove, J., Sullivan, E., Blyth, S., Zockler, C., Stanwell-Smith, D., Kingston, N., Martin, C., Spalding, M., Fletcher, S., 2017. A global map of saltmarshes. Biodivers. Data J. 5, e11764. Mitsch, W.J., Gosselink, J.G., 2000. Wetlands. third ed. John Wiley & Sons, Inc., USA. Obrist, D., Kirk, J.L., Zhang, L., Sunderland, E.M., Jiskra, M., Selin, N.E., 2018. A review of global environmental mercury processes in response to human and natural perturbations: changes of emissions, climate, and land use. Ambio 47, 116–140. Odum, E.P., 1971. Fundamentals of Ecology. third ed. W.B. Saunders Company, Philadelphia-London-Toronto. Pacyna, E.G., Pacyna, J.M., Pirrone, M., 2001. European emissions of atmospheric mercury from anthropogenic sources in 1995. Atmos. Environ. 35 (17), 2987–2996. Patra, M., Bhowmik, N., Bandopadhyay, B., Sharma, A., 2004. Comparison of mercury, lead and arsenic with respect to genotoxic effects on plant systems and the development of genetic tolerance. Environ. Exp. Bot. 52, 199–223. Pedro, S., Duarte, B., Almeira, P.R., Caçador, I., 2015. Metal speciation in salt marsh sediments: influence of halophyte vegetation in salt marshes with different morphology. Estuar. Coast. Shelf Sci. 167, 248–255. Pendleton, L., Donato, D.C., Murray, B.C., Crooks, S., Jenkins, W.A., Sifleet, S., Craft, C., Fourqurean, J.W., Kauffman, J.B., Marbà, N., Megonigal, P., Pidgeon, E., Herr, D., Gordon, D., Baldera, A., 2012. Estimating global “blue carbon” emissions from conversion and degradation of vegetated coastal ecosystems. PLOS One 7 (9), e43542. Peverly, J.H., Surface, J.M., Wang, T., 1995. Growth and trace metal absorption by Phragmites australis in wetlands constructed for landfill leachate treatment. Ecol. Eng. 5, 21–35. Pezeshki, S.R., 2001. Wetland plant responses to soil flooding. Environ. Exp. Bot. 46, 299–312. Pezeshki, S.R., DeLaune, R.D., 2012. Soil oxidation-reduction in wetlands and its impact on plant functioning. Biology 1, 196–221. Pietrini, F., Iannelli, M.A., Pasqualini, S., Massacci, A., 2003. Interaction of cadmium with glutathione and photosynthesis in developing leaves and chloroplasts of Phragmites australis (Cav.) Trin. ex Steudel. Plant Physiol. 133, 829–837. Psaras, G.K., Constantinidis, Th., Cotsopoulos, B., Manetas, Y., 2000. Relative abundance of nickel in the leaf epidermis of eight hyperaccumulators: evidence that the metal is excluded from both guard cells and trichomes. Ann. Bot. (Lond.) 86, 73–78. Rahn Jr., W.R., 1973. The Role of Spartina alternijlora in the Transfer of Mercury in a Salt Marsh Environment. Master of Science Thesis. Georgia Institute of Technology July 1973. Randall, P.M., Yates, B.J., Lal, V., Darlington, R., Fimmen, R., 2013. In-situ subaqueous capping of mercury-contaminated sediments in a fresh-water aquatic system, part II— evaluation of sorption materials. Environ. Res. 125, 30–40. Redjala, T., Zelko, I., Sterckeman, T., Legue, V., Lux, A., 2011. Relationship between root structure and root cadmium uptake in maize. Environ. Exp. Bot. 71 (2), 241–248.

Santos, D., Duarte, B., Caçador, I., 2014. Unveiling Zn hyperacumulation in Juncus acutus: implications on the electronic energy fluxes and on oxidative stress with emphasis on non-functional Zn-chlorophylls. J. Photochem. Photobiol. B. Biol. 140, 228–239. Santos, D., Duarte, B., Caçador, I., 2015. Biochemical and photochemical feedbacks of acute Cd toxicity in Juncus acutus seedlings: the role of non-functional Cd-chlorophylls. Estuar. Coast. Shelf Sci. 167, 228–239. Schreiber, U., Schliwa, U., Bilger, W., 1986. Continuous recording of photochemical and non-photochemical chlorophyll fluorescence quenching with a new type of modulation fluorometer. Photosynth. Res. 10 (1), 51–62. Schroder, P., Lyubenova, L., Huber, C., 2009. Do heavy metals and metalloids influence the detoxification of organic xenobiotics in plants? Environ. Sci. Pollut. Res. 16, 795–804. Shimazaki, K., Doi, M., Assmann, S.M., Kinoshita, T., 2007. Light regulation of stomatal movement. Annu. Rev. Plant Biol. 58, 219–247. Sizmur, T., McArthur, G., Risk, D., Tordon, R., O'Driscoll, N.J., 2017. Gaseous mercury flux from salt marshes is mediated by solar radiation and temperature. Atmos. Environ. 153, 117–125. Skinner, K., Wright, N., Porter-Goff, E., 2007. Mercury uptake and accumulation by four species of aquatic plants. Environ. Pollut. 14, 234–237. Smith, L.M., Reinfelder, J.R., 2009. Mercury volatilization from salt marsh sediments. J. Geophys. Res. 114, G00C09. Sonke, J.E., Heimbürger, L.E., Dommergue, A., 2013. Mercury biogeochemistry: paradigm shifts, outstanding issues and research needs. Compt. Rendus Geosci. 345 (5–6), 213–224. Sousa, A.I., Caçador, I., Lillebø, A.I., Pardal, M.A., 2008. Heavy metal accumulation in Halimione portulacoides: intra- and extra-cellular metal binding sites. Chemosphere 70, 850–857. Suntornvongsagul, K., Burke, D., Hamerlynck, E., Hahn, D., 2007. Fate and effects of heavy metals in salt marsh sediments. Environ. Pollut. 149, 79–91. Tomiyasu, T., Matsuo, T., Miyamoto, J., Imura, R., Anazawa, K., Sakamoto, H., 2005. Low level mercury uptake by plants from natural environments — mercury distribution in Solidago altissima L. Environ. Sci. 12 (4), 231–238. Urban, J., Ingwers, M.W., McGuire, M.A., Teskey, R.O., 2017. Increase in leaf temperature opens stomata and decouples net photosynthesis from stomatal conductance in Pinus taeda and Populus deltoides × nigra. J. Exp. Bot. 68 (7), 1757–1767. Válega, M., Lillebø, A.I., Caçador, I., Pereira, M.E., Duarte, A.C., Pardal, M.A., 2008a. Mercury mobility in a salt marsh colonised by Halimione portulacoides. Chemosphere 72, 1607–1613. Válega, M., Lillebø, A.I., Pereira, M.E., Caçador, I., Duarte, A.C., Pardal, M.A., 2008b. Mercury in salt marshes ecosystems: Halimione portulacoides as biomonitor. Chemosphere 73 (8), 1224–1229. Wang, Y.D., Greger, M., 2004. Clonal differences in mercury tolerance, accumulation, and distribution in willow. J. Environ. Qual. 33, 1779–1785. Weis, J.S., Weis, P., 2004. Metal uptake, transport and release by wetland plants: implications for phytoremediation and restoration. Environ. Intern. 30, 685–700. Williams, T.P., Bubb, J.M., Lester, J.N., 1994. Metal accumulation within salt marsh environments: a review. Mar. Pollut. Bull. 28, 277–290. Windham, L., Weis, J.S., Weis, P., 2001. Patterns and processes of mercury release from leaves of two dominant salt marsh macrophytes, Phragmites australis and Spartina alterniflora. Estuaries 24 (6), 787–795. Yang, X., Feng, Y., He, Z., Stoffella, P.J., 2005. Molecular mechanisms of heavy metal hyperaccumulation and phytoremediation. J. Trace Elem. Med. Biol. 18, 339–353. Zhu, X.G., Govindjee, Baker, N.R., Sturler, E.D., Ort, D.R., Long, S.P., 2005. Chlorophyll a fluorescence induction kinetics in leaves predicted from a model describing each discrete step of excitation energy and electron transfer associated with photosystem II. Planta 223, 114–133.