2. Publications in peer-reviewed journals and manuscripts
2.1. Cadmium toxicity investigated on physiological and biophysical level under
the aquatic model plant Ceratophyllum demersum L.
Elisa Andresen1, Sophie Kroenlein1, Hans-Joachim Stärk2 and Hendrik Küpper1,3
1) University of Konstanz, Department of Biology, D-78457 Konstanz, Germany
2) UFZ – Helmholtz Centre for Environmental Research, Department of Analytical Chemistry, Permoserstr. 15, D-04318 Leipzig, Germany
3) University of South Bohemia, Faculty of Biological Sciences and Institute of Physical Biology, Branišovská 31, CZ-370 05 České Budejovice, Czech Republic
Unpublished manuscript
Abstract
Cadmium is an important environmental pollutant and poisonous to most organisms. We investigated the mechanisms of Cd toxicity in the aquatic, rootless model plant Ceratophyllum demersum using environmentally relevant conditions including nature-like light and temperature cycles and Cd concentrations in the nM range. As expected, the level of inhibition increased with increasing Cd concentrations. The threshold concentration for most parameters was 20 nM, below, hardly any stress symptoms were observed. The first site of inhibition was photosynthesis (measured as Fv/Fm, ΦPS II), followed by increased production of reactive oxygen species, most likely a follow reaction of dysfunctional photosynthesis and energy dissipation. Cd treatment induced changes in pigment contents, reducing Chl and increasing quenching pigments. All observed effects were more pronounced in plants cultivated under high light conditions compared to low light conditions.
Introduction
The heavy metal cadmium is an important environmental pollutant and toxic to most organisms. Concentrations in the Earth’s crust are rather low (0.1-0.5 ppm, Maret and Moulis, 2013), but mining, smelting and industrial use (e.g. as plastic stabilzer, in pigments and in NiCd batteries) still increase the amount of Cd in the environment. Cd is highly water soluble and can be easily taken up by plants, thus entering the human food chain (McLaughlin et al., 1999). Due to chemical similarity with Zn, Cd is mainly taken up via transporters with similar affinity for Cd and Zn, and many toxic effects are related to Zn replacement or limitation (Clemens, 2006). Sewage sludge and organic fertilizers contaminated with Cd bring it directly into contact with crop plants and should be avoided for this reason (He et al., 2005; McBride et al., 1997).
Under specific circumstances, Cd was found to have a metabolic function. The first observation was that diatoms under severe Zn-limitation show enhanced growth, when Cd was added (Price and Morel, 1990). Later experiments revealed that those diatoms express a carbonic anhydrase (CA), which can have Cd in its active center, when Zn is not available (CDCA, ζ-CA). Growth enhancement under Zn-limitation was also observed in other species of marine phytoplankton, which do not possess the cdca gene (recently reviewed by Xu and Morel, 2013), suggesting another biochemical role of Cd, which could explain the micronutrient-like distribution of Cd in the oceans (Pai and Chen, 1994).
Besides the above mentioned positive function, there is overwhelming evidence for cadmium toxicity in plants (Andresen and Küpper, 2013). Cadmium can interfere with all parts of a plants’ metabolism: The uptake or translocation of other, essential nutrients can be inhibited, because Cd competes with other ions for the binding sites (Dong et al., 2006), making deficiency or imbalance of essential nutrients a part of Cd toxicity. Not only the binding sites of transporters can be filled by Cd, other ions in enzymes can be replaced by Cd as well. The substitution in a structural site can, but also may not result in great changes in the protein structure. But if the substitution occurs in the active center, it will make the enzyme inoperable, as redoxreactions are not possible with redox-inert Cd (Maret and Moulis, 2013).
A great threat for all photosynthetic organisms is the inhibition of photosynthesis. It was shown that Cd treatment led to decreased photosynthetic activity in various plant species (see reviews by Andresen and Küpper, 2013; Küpper and Kroneck, 2005) with a much stronger inhibition of PS II compared to PS I (Atal et al., 1991; Clijster and Van Assche, 1985). The replacement of the Mg2+-ion in the chlorophylls of the reaction centers and in the light harvesting complexes (LHCs) is one result of heavy-metal treatment in photosynthetic
organisms. The heavy-metal substituted chlorophylls ([hms]-Chls) are unsuitable for photosynthesis (Kowalewska et al., 1992, 1987, Kowalewska and Hoffmann, 1989). The mode of substitution is strongly dependent on the irradiance as shown by Küpper et al. (1998, 2002), leading to stronger inhibition of photosynthesis by heavy metals in high light (Cedeno-Maldonado and Swader, 1972). Cadmium can enhance the presence of reactive oxygen species (ROS) in plants and algae (reviewed by Andresen and Küpper, 2013; Sandalio et al., 2009; Pinto, 2003). As Cd is a redox-inert metal, ROS cannot be produced directly via Fenton or Haber Weiss reaction (Wardman and Candeias, 1996), but indirectly by impairment of respiration and photosynthesis leading to the mis-transfer of electrons to oxygen instead of their target molecule. Alternatively, the antioxidative system can be inhibited by Cd, resulting in a reduced removal of existing ROS (Sandalio et al., 2001).
All of these effects have been observed in plants, but in most studies, very high, environmentally unrealistic Cd concentrations (upto mM, e.g. Sigfridsson et al., 2004) were used. Cd concentrations in unpolluted environments range from 0.2-0.4 nM (e.g. Lake Constance: Zweckverband Bodensee-Wasserversorgung, 2011; Petri, 2006) to 5 nM in slightly polluted rivers in Germany (Bachor et al., 2012). Even in a heavily polluted stream in Nigeria, the highest concentration measured was 1.3 µM (Ahmed et al., 2011). Effects observed under very high Cd concentrations are therefore questionable as plants would never be exposed to them in nature. The effects may be very different under acute (hours to days) vs. chronic (days to months) toxicity. Additionally, laboratory conditions simulating natural conditions (e.g. light and temperature cycles apart from switch on / switch off) may also lead to new insights into toxicity mechanisms. Many studies show that Cd is toxic to plants, that photosynthesis is inhibited, or that Cd treatment induces reactive oxygen species, but the causal relationship often remained unclear.
This study aimed to fill this gap and investigate the biochemical and biophysical mechanisms of Cd toxicity under environmentally relevant conditions. We used nature-like light and temperature conditions and applied Cd concentrations from less than 1 nM to 200 nM.
Material and methods
Plant material and culture conditions
The rootless, submerged and free-floating macrophyte Ceratophyllum demersum L. was used for the stress experiments. The strain was obtained from an aquaria shop and continuously cultivated since 2005 in hydroponic solution with 12 h day/12 h night light conditions provided by two Osram FLUORA® fluorescent and two warm white fluorescent tubes (Osram, München, Germany) and a temperature cycle from 18°C at 6 a.m., over 20°C at 9 a.m., to a maximum of 22°C at 3 p.m., back over 20°C at 9 p.m. to 18°C again at 6 a.m. The nutrient solution (SMNS, submerged macrophyte nutrient solution) was optimized for growth of submerged macrophytes and resembled the situation of typical oligotrophic waters, in particular soft waters (Andresen et al., 2013b). The pH was adjusted to 7.8 with KOH. All experiments were carried out under simulations of natural light and temperature conditions:
12 h night and 12 h sinusoidal light cycle with maximal irradiances at 500 µmol photons m-2 s-1 (supplied by Dulux L 55 W / 12_954, OSRAM München, Germany) for high light experiments and 60 µmol photons m-2 s-1 for low light experiments. The temperature cycle was 19°C at 6 a.m., 21.5°C at 9 a.m., 24°C at 3 p.m., 23 C at 9 p.m., 19 C at 6 a.m. For each treatment around 1.5 g (fresh weight) of plants were used. The number of individual plants was consistent for each concentration within the same experiment. Differences occurred due to weight and size of the plants at treatment start. Each aquarium contained 2 L of medium to secure a low biomass to water volume ratio, which was also constantly aerated by room air. The nutrient solution was continuously exchanged (flow rate 0.5 L day-1) to ensure that the metal uptake into the plants was limited only by the concentration, but not by the amount of nutrient solution available. After at least one week of acclimation to the experimental light conditions, cadmium was applied as CdSO4 to the medium. The concentrations were 0.2, (background, no Cd added), 0.5 (LL only), 1, 2, 5, 10, 20, 50, 100, and 200 nM. Epiphytic algae and cyanobacteria were weekly removed from the macrophytes by gentle brushing in SMNS (without micronutrients) and the aquaria cleaned with ddH2O.
Three experiments were carried out under low light (LL) conditions, two under high light (HL) conditions. A third HL experiment was performed, in which the plants showed the same trends of inhibitions, until they died after 5 weeks for unknown reasons.
All of the described measurements of physiological parameters were done using the nutrient solution SMNS without micronutrients (Rochetta and Küpper, 2009).
Fluorescence kinetic microscopy
Physiological changes in the plants induced by cadmium treatment were determined by two-dimensional (imaging) microscopic measurements using the Chl fluorescence kinetic microscope (Küpper et al., 2000a, 2007a). One leaf from the 5th nodium counted from the apex of the plant was fixed in the measuring chamber by gas-permeable cellophane and the area just before the branching (approximate size of 1.1 x 1.1 mm) used for the spatially and spectrally resolved kinetic measurements. A continuous flow of tempered (25°C) culture medium kept the sample under physiological conditions. A detailed description of the microscope and the used protocols of the Kautsky induction can be found in Küpper et al.
(2007a) and Andresen et al. (2010). These measurements were done weekly, one day after the cleaning. The leaves from the FKM measurement were frozen in liquid nitrogen and stored at -80°C after use for pigment analyses.
Oxygen exchange
After 6 weeks of treatment net photosynthetic oxygen release and respiratory oxygen uptake were measured with a Clark-type electrode (CellOx® 325, OxiCal-SL, WTW, Weilheim, Germany) in a custom-made 200 ml measuring chamber at 25°C. Oxygen uptake was measured in the dark, and oxygen release under increasing light intensities (from darkness to 466 µmol photons m-2 s-1). Data were recorded and analyzed using the OxyCorder measuring device with the software Oxywin 3.1 (Photon Systems Instruments, Brno, Czech Republic).
Determination of superoxide
Superoxide was determined with the dye MCLA (2-Metyl-6-(4-Methoxyphenyl)-3,7-Dihydroimidazol[1,2-A]pyrazin-3-One, Invitrogen). Two leaves from the 5th nodium were incubated in 1990 µl SMNS and illuminated for 30 min at 26 µmol photons m-2 s-1. The leaves were removed and 10 µl 100 µM MCLA were added to the medium to achieve a concentration of 50 nM. One molecule MCLA bound to one molecule O2
and generated one photon, which was detected using a luminometre (LKB WALLAC 1250 + KEITHLEY 177) and the software Oxywin 3.1. Superoxide in medium incubated without leaves and the baseline (blank value, no sample) were also measured. The relative superoxide production was defined as (value from leaves - value from medium) / (value from medium - baseline).
The leaves from the superoxide measurement were frozen in liquid nitrogen and stored at -80°C after use for pigment analyses.
Determination of H2O2 production with PF2
Intracellular hydrogen peroxide (H2O2) production was determined with a H2O2-specific fluorescent indicator based upon a boronate deprotection mechanism (Chang et al., 2004;
Lippert et al., 2011), Peroxyfluor-2 (PF2, Dickinson et al., 2010). Two leaves from the 5th nodium counted from the apex of the plant were incubated in 100 µM PF2 in 0.5 ml of SMNS for 30 min in the dark. After 30 min of destaining in 15 ml SMNS in the dark, the leaves were fixed to the measuring chamber of the FKM. The medium was exchanged after every measurement and all tubes were washed with ddH2O. The leaves were frozen afterwards in liquid nitrogen and stored at -80°C until pigment analyses. The H2O2-specific fluorescence was measured in the FKM using the filter set from AHF (Tübingen, Germany) with an excitation filter 420-500 nm (AHF F42-468), dichroic mirror 505 nm (AHF F71-302) and 520-550 nm emission filter (AHF F47-535). Flashes of blue supersaturating light were given at increasing signal integration times (20 µs up to 20 ms). For each sample, the integration time that led to the highest signal intensity without saturating the camera was chosen for quantitative analysis to avoid noise at too short exposure times, and oversaturation of the camera at too long exposure times. For each exposure time, hundreds of single pictures were taken and averaged. Background signal for each exposure time was subtracted automatically via a measurement without light.
Images of the measurement were analyzed with the FKM software and the fluorescent signal was re-calculated to one integration time according to an empirical calibration of exposure times vs. signal intensity.
Harvest
After 6 weeks of treatment, the plants were washed in SMNS and young and old tissues were separated from each other. Young tissues being 4 cm from the apex and 2 cm from the apex of each branch, old tissues 8 cm from the stem end and the rest of the branches. Remaining SMNS was removed by shaking, and the plants were frozen in acid washed tubes in liquid nitrogen. Samples were stored at -80°C until further analysis.
Determination of pigment content
Harvested material and weekly frozen leaves were lyophilized and ground with sand and a few grains of Bis-Tris (Sigma-Aldrich, St Louis, MO, USA). Pigments were extracted in 1 ml 100% acetone over night at 4°C in the dark. Spectra of pigment extracts were measured with the UV/VIS/NIR absorption spectrophotometer Lambda750 (Perkin-Elmer, Waltham, MA, USA) at a spectral bandwidth of 0.5 nm, with 0.5 nm sampling interval and from 330 to 750 nm. Pigment composition was analyzed using the Gauss-Peak-Spectra method by Küpper et al. (2000b, 2007b).
Determination of starch accumulation
The starch extraction was based on the Total Starch Assay from Megazyme (AOAC Method 996.11 and ACCC Method 76.13, Megazyme, Wicklow, Ireland). The recommended protocol including pre-dissolving by KOH was downscaled for the analyses of starch in Ceratophyllum demersum (Andresen et al., 2013b; Thomas et al., 2013).
Determination of metal accumulation
The lyophilized plant material was acid digested following the protocol by Zhao et al. (1994).
All glassware used had previously been acid washed (5% HNO3). Acids for the digestion were of supra or ultrapure quality (ROTIPURAN, Roth, Karlsruhe, Germany). Analyses were done with a graphite furnace atomic absorption spectrometer (GBC 932 AA with GF 3000, Breaside, VIC, Australia). Standard solutions for Cd, Cu, Fe and Zn were diluted from AAS Standards (TraceCERT, Sigma-Aldrich, St. Louis, MO, USA). The digested plant samples were appropriately diluted to optimal detection range with suprapure (for Cd, Cu and Fe) or ultrapure (for Zn) 1.66% HCl.
Determination of elements in the solutions
Element concentrations in the nutrient solution were determined in the barrels and in the aquaria before the plants were harvested, i.e. after one week of treatment. The respective concentrations were analyzed using an inductively coupled plasma sector field mass spectrometer ICP-sfMS (Element XR, Thermo Fisher Scientific, Waltham, MA, USA). Prior to analysis ICP-sfMS parameters were optimized every day and samples were diluted 1:20
prior to analysis. The calibration with up to 5 standards was verified using the following reference materials: SLRS-5 (River Water Reference Material for Trace Metals, NRCC), SPS-SW1 (Surface Water Level 1, Spectra Pure Standards) and SRM 1643e (Trace Elements in Water, NIST). The instrumental parameters for determination of the investigated elements are reported in Andresen et al., 2013b.
Statistics
Two-way and three-way analysis of variance (ANOVA) was done in SigmaPlot 11 (SPSS Science, USA) at the significant level of P<0.05 with Cd concentration, weeks of treatment for the weekly measured data and Cd concentration, age and light condition for the data obtained from the harvested material. The Holm-Sidak method was used for multiple comparison procedure.
Results
Growth and visible symptoms of Cd toxicity
The plants treated with 200 nM of Cd under LL conditions stopped growth directly after start of the treatments. Growth of the meristems of these plants was retarded and the meristems looked compressed. Growth decreased further until the plants had to be harvested after 4 weeks (Fig. 1). From the second week on, growth stop occurred also in the second and third highest Cd concentration (50 and 100 nM). The threshold concentration of growth inhibition and visible impairment was 20 nM Cd. Under HL conditions, some growth inhibition was also observed in the low Cd concentrations and the control due to excessive growth of epiphytic algae and cyanobacteria. The sequence of visible toxicity symptoms was slightly different for the two light conditions. Under HL conditions, the visible changes of the meristem and a reduction in the internodial growth occurred for the three highest Cd concentrations already in the first week. Between week 3 and 4, the plants treated with 20 nM Cd were also affected in plants from LL and HL conditions. All plants treated with 10 nM or less Cd did not show visible symptoms of toxicity.
Photosynthetic parameters
Cadmium treatment led to a diminished Fm under HL conditions from the first week of treatment (200 nM; Fig. 2) with subsequent reductions until 20 nM Cd in the fourth week (P<0.001). The reduction in Fm was consistent with a decreased Chl concentration with similar pattern. Under LL conditions, the effect was not as pronounced, but significant for the plants treated with 100 nM Cd.
Cadmium treatment led to a diminished photochemical efficiency of PS II as measured by Fv/Fm (Fig. 2). From the second week of treatment on, there was a distinct reduction observable: Plants treated with 50 nM, 100 nM or 200 nM Cd had decreased values, while no effect was observed below 50 nM (P<0.001). From the third week on, the plants treated with 100 nM and 200 nM Cd yielded values reduced by 50% compared to the treatment start and the control samples. The reduction in Fv/Fm was much more apparent in plants from the HL experiments compared to LL. A noticeable decrease was only observed in the macrophytes treated with 100 nM and 200 nM from week 3 on.
Photochemical fluorescence quenching displays the operating efficiency of PS II (ΦPS II, or Fv’/Fm’). The same trend as for Fv/Fm was visible for ΦPS II in HL at the beginning of the actinic light phase (i1), after acclimation in actinic light (i6), and during recovery in darkness after actinic light (r1 and r5; Fig. 3). The decrease was observed from the second week on for all plants cultivated with 50 nM or more for all 4 parameters. A different behavior was observed in the LL experiment. Photochemical quenching in actinic light was reduced in all plants of Cd treatment with 50 nM or higher from the first weeks of treatment on (Fig. 3). In the last weeks, also the plants treated with 20 nM showed inhibition. In the dark phase after actinic light (r1), a strong decrease was observed only for the plants treated with the highest Cd concentration. After approximately 200 s in darkness (r5), hardly any effect was observable, besides a reduction for the three highest concentrations (P<0.001). The other plants were able to relax after the actinic light phase.
Unlike the photochemical quenching, there was no clear trend detectable in the non-photochemical quenching (NPQ, Fig. 4) of the HL and LL plants during actinic light (i). At the end of the relaxation period (r5), however, the plants treated with 10-50 nM Cd had enhanced NPQ from the third and fourth week on, which was also observable at the spectrally resolved level (Fig.5). An enhanced quenching occurred at two positions, at approximately 690 nm and 750 nm in the plants treated with 20-100 nM towards the end of the treatment (week 5+6).
Oxygen exchange
After six weeks of treatment, oxygen exchange was measured for plants in darkness and in increasing light intensities. Net oxygen evolution increased with increasing irradiances. Cd treatment led to delayed oxygen evolution in HL plants, most prominently already from 2 nM onwards, and LL plants from 20 nM Cd (Fig. 6). Measurement of oxygen release in low irradiance led to higher amounts of O2 in plants grown under HL conditions, compared to LL grown plants. Although LL grown plants have bigger antenna systems, they have comparably fewer RC than HL grown plants (Taiz and Zeiger, 2007) and therefore less oxygen evolution.
Respiration showed no clear trend under HL conditions (measured after 6 weeks of treatment) suggesting that the respiratory electron chain in mitochondria was not so much affected by Cd treatment compared to photosynthesis. Under LL conditions, respiration after actinic light was increased in the highest measureable concentration (50 nM). Generally, oxygen uptake was higher in the plants of the HL conditions (P<0.001).
Production of reactive oxygen species
Cadmium treatment enhanced the release of superoxide (O2
•-) from the leaves into the medium (Fig. 7, upper part). The relative production of superoxide was higher and started earlier (from second week on) from those leaves of the HL experiment. Despite some noise, the threshold concentration for enhanced O2•- under HL conditions was 10 nM Cd, although significant differences were obtained between the 50 nM and all plants from 0.2 nM-10 nM.
In the leaves from LL, an increase was only observed in the three highest Cd concentrations in the beginning and towards the end of the treatment (week 5-6; Fig. 7, upper part).
Superoxide production in those three treatments was different from the control treatment (P<0.001).
Hydrogen peroxide (H2O2) was elevated in response to Cd treatment in HL plants
Hydrogen peroxide (H2O2) was elevated in response to Cd treatment in HL plants