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Keywords:

  • flow cytometry;
  • C11-BODIPY581/591;
  • oxidative stress;
  • lipid oxidation;
  • Chlamydomonas reinhardtii;
  • metals;
  • nanoparticles;
  • diethyldithiocarbamate;
  • diuron

Abstract

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Lipid oxidation is a recognized end point for the study of oxidative stress and is an important parameter to describe the mode of micropollutant action on aquatic microorganisms. Therefore, the development of quick and reliable methodologies probing the oxidative stress and damage in living cells is highly sought. In the present proof-of-concept work, we examined the potential of the fluorescent dye C11-BODIPY591/581 to probe lipid oxidation in the green microalga Chlamydomonas reinhardtii. C11-BODIPY591/581 staining was combined with flow cytometry measurements to obtain multiparameter information on cellular features and oxidative stress damage within single cells. First, staining conditions were optimized by exploring the capability of the dye to stain algal cells under increasing cell and dye concentrations and different staining procedures. Then lipid oxidation in algae induced by short- and long-term exposures to the three metallic micropollutants, copper, mercury, and nanoparticulate copper oxide, and the two organic contaminants, diethyldithiocarbamate (DDC) and diuron was determined. In this work we pointed out C11-BODIPY591/581 applicability in a wide range of exposure conditions, including studies of oxidation as a function of time and that it is suitable for in vivo measurements of lipid oxidation due to its high permeation and stability in cells and its low interference with algal autofluorescence. © 2013 International Society for Advancement of Cytometry

The measurement of induced oxidative stress and associated cellular damage in aquatic microorganisms has become, in the last years, a recognized end point and an accepted paradigm to assess and compare the toxicity of different micropollutants on microalgae [1]. Consequently, the measurement of oxidative stress and associated cellular damage, together with standard ecotoxicological endpoints such as growth and photosynthesis inhibition are of high relevance for understanding the mode of micropollutant action. Toxic metals, nanoparticles, and organic compounds [2-5], and also some physical stressors like UV radiation [6] have the potential to cause oxidative damage in microalgae. The methods currently used to assess oxidative stress are numerous and range from direct quantification of reactive oxygen species (ROS) to the indirect evaluation of oxidative stress through measurement of the induced cellular effects [7]. In studies focused on the evaluation of oxidative stress in algae, these methods are currently applied and are either based on the evaluation of cellular damage (e.g., lipid [8, 9] and protein oxidation [10]) or on the quantification of antioxidant enzymes and radical scavengers [11-13]. In recent years, the use of fluorescent probes specifically designed to detect ROS in vivo or to label ROS-induced cellular damage has notably increased since these methods couple high sensitivity with simple use [14]. Among the fluorescent probes available for the investigation of oxidative stress in living cells, probes based on dihydrofluoresceine diacetate have been applied with microalgal cells [15, 16]. However, their use is limited by their high photosensitivity, low dye penetration in the cells and the necessity to add substances to enhance membrane permeability [17, 18]. To our knowledge there are no dyes that are systematically applied for the detection of lipid oxidation in microalgal cells, which is why in this work we propose a lipophilic fluorescent dye 4,4-difluoro-5-(4-phenyl-1,3-butadienyl)-4-bora-3a,4a-diaza-s-indacene-3-undecanoic acid (C11-BODIPY581/591) to probe oxyl-radical induced lipid oxidation by flow cytometry (FCM). C11-BODIPY581/591 is a fatty acid analog with specific fluorescent properties [19]. When excited with blue light at 488 nm wavelength, the molecule has a constitutive fluorescence emission with a maximum at 595 nm. Following oxyl-radical induced oxidation, the fluorescence emission shifts to shorter wavelengths with a maximum emission at 520 nm. Due to its lipophilic properties, the molecules easily enter the lipid bilayer and once inside the cellular membrane are subject to oxidation by oxyl-radicals together with the endogenous fatty acids [14]. C11-BODIPY581/591 is a good candidate for routine analyses and ecotoxicity tests; it has a high quantum yield of fluorescence emission that assures a good signal measurement even at low concentration, and it has high photostability and its fluorescence emission is not prone to pH changes and solvent polarity. Moreover, its oxidation induced fluorescence shift can be triggered only by oxy-radicals and not by superoxides, nitric oxides, transition metal ions and hydroperoxides [19]. The last feature is particularly important in ecotoxicity testing since the specificity of the dye for radical species reduces the occurrence of false positives generated by the reaction of the probe with other chemicals present in the medium. C11-BODIPY581/591 has been already proposed to determine lipid peroxidation [20] and has been used as a lipid peroxidation probe in mammalian cells [21-25]. Furthermore, comparison of lipid peroxidation obtained by using C11-BODIPY581/591 with mass spectrometry quantification of an oxidized lipid, the 1-stearoyl-2-arachidonyl-sn-glycero-3-phosphocholine, revealed the tendency of C11-BODIPY581/591 to overestimate lipid peroxidation in the cells but confirmed its capability to highlight oxidative stress conditions at the membrane level that have the potential to cause lipid peroxidation [26]. Dyes from the BODIPY-family have been also applied in studies on microalgae, notably in algal biotechnology for the evaluation of intracellular lipids [27] and the estimation of the antioxidant activity of algal pigments [28]. Recently, the potential of this dye was explored to study lipid peroxidation induced by silver nanoparticles on the microalgae Chlorella vulgaris and Dunaliella tertiolecta [15]. In this study, the results obtained with C11-BODIPY581/591 positively correlated with the measurement of cellular oxidative stress determined by a fluorescein diacetate probe. However, no information was reported concerning the staining procedure. To our knowledge, no other studies with microalgae were published using this dye in an ecotoxicological context. Therefore, there is a lack of systematic knowledge about the influence of different staining conditions on the dye performance in microalgal species, as well as no established procedures.

FCM is a single cell technique that provides a multi-parameter assessment of the physiological state of individual cells of a population. In studies concerning algae, this technique has been extensively applied to investigate the distribution of phytoplankton in natural environments [26] and to discriminate, identify and isolate algal species [27, 28]. Because of its capability to discriminate different morphological features in cells of the same population, this technique is increasingly applied in the optimization of biotechnological microalgal processes [29] and in ecotoxicological studies [17, 30-34]. A combination of FCM and appropriate fluorescent dyes is used in ecotoxicology to investigate the effects induced either by metallic or organic micropollutants. Examples include testing altered membrane permeability with the propidium iodide staining [30, 31], esterase activity by FDA staining [32-34] and oxidative stress using different probes like 5-(and-6)-carboxy-2′,7′-dihydrodifluorofluorescein di-acetate (H2DFFDA) [17], 2′,7′-dichlorodihydro-fluorescein diacetate (H2DCFDA) [34] or dihydrorhodamine 123 (DHR123) [31, 33]. Nonetheless, the potential of FCM in the field of ecotoxicology is still underexploited.

The aim of the present proof-of-concept study is therefore to examine the suitability of the dye C11-BODIPY581/591 in combination with FCM for the assessment of lipid oxidation in the microalga Chlamydomonas reinhardtii under a variety of exposure conditions to different micropollutants. Microalgae were chosen as model organisms because of their important ecological functions being the base of the food chain as well as because of their wide use in ecotoxicity testing. In the adopted experimental strategy, the dye was first tested treating C. reinhardtii cells with different H2O2 concentrations and exposure durations to verify its ability to reveal oxyl-radical induced lipid oxidation and to set optimal staining conditions. Then, the lipid oxidation potential of different environmental micropollutants such as copper and mercury, copper oxide nanoparticles (CuO-NPs), and two pesticides, a generic antimicrobial agent diethyldithiocarbamate (DDC) and the herbicide diuron were quantified in a long- and short-term exposure using the optimized staining conditions.

Material and Methods

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Reagents and Glassware

All stock solutions and media were prepared in MilliQ water (>18.2 Ω MilliQ Direct system, Merck Millipore, Darmstadt, Germany) and all reagents used were of analytical grade. Stock standard solutions of 0.1 mol L–1 copper sulfate and 1,000 mg L–1 mercury in 12% nitric acid, sodium diethyldithiocarbamate trihydrate and diuron were obtained from Sigma-Aldrich (Buchs, Switzerland). Stock solutions of DDC and diuron, with concentration of 2 × 10−3M and 10–3M, respectively, were prepared in experimental medium and then filtered through 0.45 µm-pore size regenerated cellulose filters (Millipore, Billerica, USA). CuO-NPs of 99+ % purity and with a primary size of 30–50 nm were obtained from Nanostructured and Amorphous Materials (Houston, TX). CuO-NPs powder was dispersed in MilliQ water at a concentration of 2 g L–1. Prior use the stock solution was sonicated at the power of 130 W, 20 kHz for 1 min at 100% amplitude (Sonics Vibra Cell, Sonic and Materials, Newtown, USA).

All the glassware was soaked for at least 24 h in 5% v/v HNO3, rinsed three times with MilliQ water and autoclaved prior use. All the manipulation of algal cultures and liquid media were performed in sterile conditions.

Test Micro-Organism and Culture Conditions

The unicellular green alga Chlamydomonas reinhardtii strain CPCC11 was obtained from the Canadian Phycological Culture Center (CPCC, Department of Biology, University of Waterloo, Canada). C. reinhardtii was axenically cultured under mixotrophic conditions in four times diluted Tris-Acetate-Phosphate (4× diluted TAP) liquid medium [35]. Algae were grown in 150 mL glass flasks in a specialized incubator (Infors, Bottmingen, Switzerland), under rotary shaking of 100 rpm, temperature of 20°C and illumination of 110 µmol phot m–2 s–1. At mid exponential growth phase, reached after 64 h of incubation, cells where collected by gentle centrifugation (2,083g for 3 min), rinsed with and re-suspended in experimental medium. All experiments were performed in an exposure medium containing 10–4 M KH2PO4; 1.5 × 10–4 M K2HPO4; 1.7 × 10–3 M NH4NO3; 8.5 × 10–5 M CaCl2·2H2O; 10–4 M MgSO4·7H2O in 2-(N-morpholino)ethanesulfonic acid (MES, Sigma-Alrdich, Buchs, Switzerland) with a final concentration of 10–3 M and pH 6.8. No trace metals were added. There was no algal growth difference during 48 h incubation in the exposure medium and 4× diluted TAP medium used for algal growth. If not specified in the text, the initial algal density was 106 cell mL–1.

Optimization of Cell Staining with C11-BODIPY581/591

The stock solution of C11-BODIPY581/591 (Life technologies-Invitrogen, Carlsbad, USA) was prepared by dissolving 1 mg of the product in DMSO to a concentration of 1 mM. Potential interference of dye fluorescence with algal autofluorescence was evaluated by comparing the signals in FCM green, orange and red fluorescence channels before and after staining. The effect of the C11-BODIPY581/591 staining on C. reinhardtii was also explored by determination of the influence of different dye concentrations and staining times on cell size, granularity and chlorophyll autofluorescence. Controls with the dye solvent, DMSO, were also performed.

Optimization experiments were performed with algae exposed for 30 min to 5 × 10–3 M H2O2, which is a well-known oxidant inducing oxidative stress and triggering oxyl-radical formation. To determine the optimal C11-BODIPY581/591 concentration, 1.25, 2.5, and 5 µM concentrations were used. Based on the available literature [19, 20] and preliminary results staining durations of 30 min and 60 min were tested. Algal densities of 104, 105, and 106 cells mL–1 were examined keeping a constant concentration of hydrogen peroxide of 5 × 10–3 M. The long term stability of C11-BODIPY581/591 was tested by measuring the green fluorescence signal after 48 h. To verify if the dye specifically responds to intracellular oxyl-radicals and if extracellular interactions occur, the effect of light irradiance composed of 207 Wm–2 visible, 9 Wm–2 UVA, and 1.9 Wm–2 UVB, and various staining procedures were investigated. Two different series of staining experiments were performed: (i) algae were exposed to 5 × 10–3 M H2O2 for 30 min, then washed with experimental medium and stained with 2.5 µM C11-BODIPY581/591 for 30 min; (ii) algae were exposed to a mixture of 2.5 µM C11-BODIPY581/591 and 5 × 10–3 M H2O2 for 30 min and directly analyzed without intermediate washing steps. Moreover, to verify if BODIPY-stained cells reflect the dead rather than stressed cells, algal cells were first inactivated by incubation at 90°C for 60 min, and then stained with the dye. Finally, lipid oxidation was evaluated by exposing the algal cells to increasing H2O2 concentrations from 10–3 to 10–2 M and exposure times from 10 to 60 min.

Determination of C11-BODIPY581/591 Distribution in Algae

The cellular compartmentalization of the oxidized C11-BODIPY581/591 was monitored by tracking its green fluorescence with a fluorescence microscope (BX61, OLYMPUS, Volketswil, Switzerland) equipped with the fluorescent filter cube U-MWIBA3 (excitation and emission band pass of 460–495 and 510–550, respectively) after 30 min exposure to UV radiation or incubation with 5 × 10–3 M H2O2. Images of the stained cells were taken using a digital camera (XC30, OLYMPUS) and processed by Cellsens software (Cellsens dimension, OLYMPUS).

Determination of Lipid Oxidation in Cells Exposed to Different Micropollutants

To explore the potential of different micropollutants to induce lipid oxidation in algal cells, C. reinhardti was exposed for 48 h to two different concentrations of pollutants; 10–7 M and 10–5 M for Cu(II) corresponding to 6 × 10–8 M and 6 × 10–6 M free copper ion concentration ([Cu2+]), 5 × 10–7 M and 5 × 10–6 M for Hg(II), 1 and 10 mg L–1 for CuO-NPs, 10–8 M and 10–7 M for DDC and diuron. The concentrations of Cu(II), Hg(II), CuO-NPs and DDC were chosen on the basis of preliminary tests (data not shown), whereas the concentrations of diuron was chosen from the literature results for C. reinhardtii [18]. Exposure experiments were performed under the same conditions as algal growth: rotary shaking of 100 rpm, temperature of 20°C and illumination of 110 µmol phot m–2 s–1 and an initial algal density of 106 cell mL–1. In the absence of pollutants the algal growth rate in the test media was 1.41 d–1. Aliquots of 2 mL of culture were sampled after 1, 24, and 48 h incubation, then supplemented with 2.5 µM C11-BODIPY581/591, incubated for 30 min and analyzed using FCM. C11-BODIPY581/591 staining was performed in parallel on unwashed and washed culture aliquots, in order to compare the possible interference of the toxicant present in the medium with the fluorescent probe. For each condition at least three replicates were performed. Unexposed algal cultures were used as negative controls, while cells incubated for 30 min with 5 × 10–3M H2O2 were used as positive controls. Prior the addition of algae to the exposure medium, the Zetasizer determined hydrodynamic size and zeta potential of CuO-NPs were 130 ± 15 nm and −31 ± 6 mV, respectively. CuO-NPs suspensions contained 30% dissolved Cu determined by inductively coupled plasma mass spectrometry after dialysis (1,000 Da cut-off of the membrane).

Determination of Time Course of Lipid Oxidation Upon Micropollutant Exposure

The time course of lipid oxidation was determined in short term exposure experiments. To this end 106 cells mL–1 of C. reinhardtii were supplemented with 2.5 µM C11-BODIPY581/591 and incubated for 30 min. Algae pre-loaded with the dye were exposed to the same concentrations of micropollutants as in the long term experiments. The percentage of green- C11-BODIPY581/591 stained cells was measured by FCM every 10 min for a period of 90 min. For each test condition at least three independent replicates were performed.

Flow Cytometry Measurements

FCM analyses were performed using a BD Accuri C6 flow cytometer with Accuri CSampler and BD Accuri C6 Software 264.15 for data acquisition and analysis (BD Biosciences, San Jose, USA). This FCM is equipped with a 488 nm argon laser and three fluorescence detectors green 530 ± 15 nm, orange 585 ± 20 nm, and red 670 ± 25 nm. Multi-parameter information on C. reinhardtii population under different exposure conditions was obtained: (i) cell size and granularity from the forward scatter (FSC) and side scatter (SSC) measurements, respectively; (ii) chlorophyll autofluorescence from red channel; and (iii) C11-BODIPY581/591 green fluorescence from green channel. Cell size, granularity and chlorophyll autofluorescence were followed in all the tests to assess the influence of the different treatments on these cellular features (Supporting Information Fig. S1). Data were collected to 20,000 events for each sample using a flow rate of 14 µL min–1, threshold was set to 20,000 on FSC-H. Absolute number of cells to account for possible formation of cellular debris and to monitor cellular growth was determined in all treatments.

FCM Data Analysis

FCM data analysis was designed to remove eventual background noise of the instrument and to select and discriminate algal cells from cellular debris (Supporting Information Fig. S2). The log FSC-H versus log FSC-A dot-plot was used first to remove cell doublets or artifacts. Then two different plots (log SSC-A versus log FSC-A plot and count versus log red fluorescence plot) were used to discriminate between algal cells and cellular debris on the base of the different size and higher chlorophyll autofluorescence of the algae in comparison with debris. The algal population in the gate designed on the log SSC-A versus log FSC-A plot was then plotted on log FSC-A versus log green fluorescence plot designed to discriminate between C11-BODIPY581/591 stained and unstained cells. The gate corresponding to the C11-BODIPY581/591 stained cells was established using the algal cultures exposed for 1 h to 5 × 10–3 M H2O2. The shift in green fluorescence was also visualized in count versus green fluorescence plots in which the distributions obtained for unexposed control and exposed C. reinhardtii were superimposed. A gate was also designed using the results obtained from analysis of heat treated cells (Supporting Information Fig. S3) designed on the values of green and red fluorescence obtained. To determine if non-cellular events (i.e., dye aggregation, CuO-NPs aggregates, debris) can occur during the C11-BODIPY581/591 treatment, FCM signals in different detectors were collected for the experimental medium supplemented with 5 μM C11-BODIPY581/591 in the absence of algae. For the experiments involving CuO-NPs additional gating was performed to discriminate between algal cells and CuO-NPs using the difference in their size (log SSC vs log FCS) and red fluorescence. The red channel was used to separate signals with low red fluorescence (CuO-NPs) and signals with higher red fluorescence (alga chlorophyll autofluorescence) (Supporting Information Fig. S4). The percentage of events in different gates is presented as an average of the three replicates with standard deviations. For comparison, the observed growth inhibition in the presence of micropollutants is provided in Supporting Information Table S1.

Results and Discussion

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Specificity of C11-BODIPY581/591 to Probe Lipid Oxidation and Cellular Distribution

Since algal cells are autofluorescent and their signature is observed in the green, orange and red channel, it was important to first verify if the fluorescent signal of oxidized and nonoxidized dye can be observed in C. reinhardtii. When stained with C11-BODIPY581/591 no changes in green, orange, and red fluorescence were observed in unexposed algal cells, while cells exposed to 5 × 10–3 M H2O2 presented a consistent shift in green fluorescence toward higher values (Fig. 1). An increase of orange fluorescence of C. reinhardtii was observed upon staining of exposed C. reinhardtii due to the spillover of the fluorescence of the oxidized molecule whose emission peak has a shoulder at higher wavelengths [19]. For this reason, the enhanced green fluorescence characteristic for oxidized C11-BODIPY581/591 rather than the changes in the ratio between oxidized and non-oxidized forms of this dye were followed by FCM.

image

Figure 1. Histograms and pictures showing the changes in cellular green (A), orange (B), and red fluorescence (C) before and after staining with C11-BODIPY581/591 in unexposed algae and exposed to 5 × 10–3 M H2O2 for 30 min. In the histograms the red curves correspond to algal fluorescence before staining, while the green curves represent the cells stained with 2.5 µM C11-BODIPY581/591 for 30 min. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

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The specificity of C11-BODIPY581/591 to respond to intracellular oxyl-radicals was investigated by comparing different staining procedures and sources of oxidative stress. Algal cultures exposed to a high intensity of visible + UV light irradiation, that has no direct interference with the dye oxidation, showed a consistent increase of green fluorescence confirming that the dye oxidation is triggered by intracellular oxyl-radicals (Supporting Information Fig. S5). Moreover, the percentage of the stained cells exposed for 30 min to a mixture of 2.5 μM C11-BODIPY581/591 and 5 × 10–3 M H2O2 was compared to those obtained in a treatment in which the cells were exposed to the same concentration of H2O2, washed and then stained. The percentage of stained cells was comparable in the two treatments, thus showing that this dye is not prone to extracellular H2O2-mediated oxidation. The above findings also suggest that C11-BODIPY581/591 exhibits high stability in presence of peroxide. Indeed, the BODIPY oxidation is known to be triggered only by oxyl-radicals such as HO, ROO, RO, and peroxynitrite, but does not react to superoxide, singlet oxygen, hydrogen peroxide etc. [14]. Temperature inactivated cells did not present an increase in green fluorescence upon staining with the dye, further confirming that C11-BODIPY581/591 probes specifically lipid oxidation and does not respond to generically dead cells.

A consistent increase of the number of cells with enhanced green fluorescence upon exposure for 30 min to 5 × 10–3 M H2O2 and in cells exposed to UV radiation (data not shown) was confirmed by fluorescence microscopy (Fig. 1). The oxidized green fluorescent dye is not equally distributed inside the algal cells. This inhomogeneous pattern of the oxidized C11-BODIPY581/591 is possibly attributable to two interrelated effects, which could not be distinguished with our experimental design: (i) the lipophilic properties of this fatty acid analog may facilitate accumulation in lipid compartments of the cell and (ii) the different rates of oxidation of the dye in different cellular compartments. Although the orange fluorescence of the non-oxidized dye could theoretically be used to obtain information about the molecule localization prior its oxidation by oxyl-radicals, the strong chlorophyll red autofluorescence of algae preclude the possibility of exploring the localization of the non-oxidized dye.

No differences in cell size, granularity and chlorophyll autofluorescence between stained and unstained algal cells were observed showing no measurable effects of the dye on the cell characteristics and no interferences with the cell autofluorescence. The above cellular characteristics were also unaffected by the presence of 0.25% v/v DMSO, which was used to dissolve the lipophilic C11-BODIPY581/591. These results agree with the finding that DMSO did not affect the growth of two other green algal species at concentrations below 1% v/v [36].

Effect of Dye Concentration and Algal Cell Density

No significant difference was observed in the percentage of the stained cells for different C11-BODIPY581/591 concentrations and cellular densities of 104, 105, and 106 cells mL–1 (Supporting Information Fig. S6), indicating that FCM in combination with C11-BODIPY581/591 allows the evaluation of lipid oxidation of the algal membranes in a wide range of cellular densities. Algal cell numbers remained unchanged and no formation of cellular debris was detected. Moreover, no formation of C11-BODIPY581/591 aggregates was detected under the studied conditions, as revealed by fluorescence microscopy and by the lack of signals in the SSC, FCS, green, orange and red fluorescence channels when the dye was incubated in presence of H2O2 and in absence of algal cells. This observation contrasts with the finding that other BODIPY dyes such as BODIPY505/515 form precipitates that exhibit strong fluorescence that interfere with algal autofluorescence [37]. However, in the latter study the BODIPY concentrations inducing the formation of dye aggregates were between 4 and 10 times higher than those used in the present study.

Effect of Staining Duration and C11-BODIPY581/591 Cellular Stability

No significant variation of the percentage of the green stained cells (Fig. 2) was found within 60 min. The green fluorescence of the C11-BODIPY581/591 stained cells was re-examined 48 h after staining. The percentage of green stained cells was similar to those obtained for 30 and 60 min staining, suggesting a high cellular stability of this dye. A residence time inside cell membranes of up to 2 h of incubation has previously been shown for this dye [19]. The above experiments were performed in light and demonstrated also that C11-BODIPY581/591 is stable to photobleaching unlike other dyes used for measuring oxidative stress via an intermediate oxidation step by intracellular esterases such as dihydrofluoresceine diacetate.

image

Figure 2. Effect of staining duration. Dot-plot of forward scatter versus green fluorescence corresponding to 2.5 μM C11-BODIPY581/591 stained cells: (A) unstained and unexposed C. reinhardtii, (B,C) unexposed algae stained for 30 min and 60 min, and (D) H2O2 treated unstained algae, (E, F) H2O2 treated algae stained for 30 min and for 60 min. Exposure to 5 × 10–3 M H2O2 for 60 min; average cell number 9.4 × 105 ± 0.6 cells mL–1. No formation of cellular debris was observed. [Color figure can be viewed in the online issue which is available at wileyonlinelibrary.com.]

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Probing Lipid Oxidation Time Course Using C11-BODIPY581/591 Stain

Following 30 min preloading of the cells with dye, the percentage of the stained cells was monitored as a function of the exposure time for three different hydrogen peroxide concentrations (Fig. 3B). The percentage of cells experiencing lipid oxidation increased in the first 20 min in a different way depending on the H2O2 concentrations in the exposure medium. The number of stained cells reached a plateau after 30 and 45 min of exposure to 5 × 10–3 M H2O2 and 10–3 M H2O2, respectively. For algae exposed to 10–2 M H2O2, a rapid increase in the level of lipid oxidation was observed within the first 20 min, followed by a decrease in the number of green stained cells, which correlated with a simultaneous decrease in cell number and an increase of cell debris (Fig. 3A), as detected in the forward and side scatter channels of FCM.

image

Figure 3. C. reinhardtii preloaded with C11-BODIPY581/591 and then exposed to H2O2. (A) Normalized cell number as a function of time. Absolute cell number was normalized by the cell number at the beginning of the test. Initial cell densities were 9.9 × 105 ± 0.5 cells mL–1, 9.5 × 105 ± 0.2 cells mL–1, 9.4 × 105 ± 0.2 cells mL–1, and 1.0 × 106 ± 0.03 cells mL–1 for unexposed algae and exposed to 10–3, 5 × 10–3 M and 10–2 M H2O2, respectively. (B) Percentage of C11-BODIPY581/591-stained algal cells as a function of exposure time. The lines represent the fit of experimental data with a four parameter log logistic model. Staining conditions: 2.5 μM C11-BODIPY581/591, 30 min. Error bars represent the standard deviation when bigger than the symbol size, N = 3.

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Obtained results demonstrated that the use of C11-BODIPY581/591 in combination with FCM offers the possibility to follow the time evolution of the number of algal cells subject to oxyl-radical-induced lipid oxidation in cell membrane and further confirms a rapid penetration and cellular stability of the C11-BODIPY581/591. Furthermore, for a given H2O2 concentration, the percentage of stained cells was unaffected by the broad range of the conditions, namely, dye concentration, staining time and algal density, which makes this dye promising for the study of lipid oxidation in algae exposed to micropollutants. The optimized set of staining conditions using 2.5 μM C11-BODIPY581/591, a staining time of 30 min and an algal density of 10−6 cell mL−1, was used to study the potential of different pollutants to induce lipid oxidation in C. reinhardtii.

Application of C11-BODIPY581/591 to Probe Lipid Oxidation in Short and Long Term Exposure to Pollutants

The potential of Cu(II), CuO-NPs, Hg(II), DDC and diuron to induce lipid oxidation in C. reinhardtii upon short term exposure and the time course of the process were explored with cells pre-loaded with C11-BODIPY581/591 (Fig. 4). No changes in the absolute cell number or formation of algal debris were observed (Supporting Information Fig. S7). Exposure to 6 × 10–6 M Cu2+ resulted in a rapid increase in the number of cells exhibiting the signature of the oxidized C11-BODIPY581/591 attaining 50 ± 2.6% of the cells in the population after 90 min. The increase in the percentage of stained cells in C. reinhardtii exposed to 5 × 10–6 M Hg(II), 10 mg L–1 CuO-NPs, and 10–7 M DDC attaining after 90 min only 7 ± 1.4, 2.2 ± 0.1, and 9.4 ± 2.3%, respectively, was much less pronounced. Among the treatments performed with the lower pollutant concentrations only 5 × 10–7 M Hg(II) induced a significant effect with respect to the unexposed control with 2.71 ± 0.27% of C11-BODIPY581/591 stained cells after 90 min. By contrast, no C11-BODIPY581/591 stained cells were determined in the algae exposed to diuron.

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Figure 4. Evolution of the percentage of C11-BODIPY581/591 stained cells as a function of time upon exposure to Cu(II) (A), Hg(II) (B), CuO-NPs (C), DDC (D), and diuron (E). Conditions: 2.5 µM C11-BODIPY581/591, algal density 106 cell mL–1. Data are shown as mean of three replicates. Error bars represent the standard deviation when bigger than the symbol size. The lines represent the fit of experimental data with a four parameter log logistic model. The concentrations of the pollutants are given as nominal values.

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In the long term exposure experiments the two concentrations of Hg(II) and Cu(II) resulted in a comparable percentage of cells experiencing lipid oxidation after 24 and 48 h (Table 1), despite the lower percentage of affected cells induced by mercury after 90 min as revealed in the short term test. Interestingly, after 48 h, the number of green stained cells exposed to 6 × 10–6 M Cu2+ was lower than that found after 24 h exposure, indicating a possible recovery of C. reinhardtii and activation of detoxification mechanisms in algae [38]. Exposure of C. reinhardtii to 10 mg L–1 CuO-NPs resulted in a significant but lower percentage of affected cells even after 48 h exposure. Only 6% and 34 ± 7.2% of the cells treated with 10–7 M diuron or DDC exhibited lipid oxidation.

Table 1. Percentage of C11-BODIPY581/591 stained Chlamydomonas reinhardtii cells upon exposure to different micropollutants for 24 and 48 h
PollutantCopperMercuryCuO-NPsDDCDiuron
  1. The concentrations are given as nominal values. For Cu(II), the concentrations are expressed as free copper ion. Species distribution for Hg(II) includes: 48% Hg(OH)2, 7.8% HgCl2 and 43% HgClOH determined with Visual MINTEQ ver. 3.0.

Conc.6 × 10–8 M6 × 10–6 M5 × 10–7 M5 × 10–6 M1 mg L–110 mg L–110–8 M10–7 M10–8 M10–7 M
 Percentage of C11-BODIPY581/591 stained cells  
24 h-exp.2.3 ± 0.799.7 ± 0.031.9 ± 0.287.3 ± 3.22.2 ± 0.95.1 ± 1.80.8 ± 0.20.9 ± 0.10.4 ± 0.20.9 ± 0.1
48 h-exp.2.1 ± 0.287.5 ± 0.61.2 ± 0.499.6 ± 0.28.4 ± 0.141 ± 11.54.3 ± 3.134 ± 7.20.6 ± 0.066.6 ± 1.3

The results of short and long term exposure experiments are consistent with available literature concerning the mode of action of pollutants. For example, copper is known to induced oxidative stress in C. reinhardtii [17]; however, little information is available concerning the potential for lipid oxidation and only for high dose treatment [39]. Similarly, no oxidative effects was observed below 10–6 M Hg(II), while above 4 × 10–6 M a consistent increase of lipid peroxidation was reported for C. reinhardtii [11]. The CuO-NPs induced oxidative damage of cellular membranes agreed with findings showing that CuO-NPs cause oxidative stress to microalgae [40]. Lower percentages of cell experiencing lipid oxidation by DDC and diuron comply with the mode of action of this organic toxicant. DDC is a superoxide dismutase (SOD) inhibitor, thus, contrarily to other toxicants, it acts on an enzyme involved in the antioxidant defense, reducing the self-protection of cells from superoxide anions and radicals [41]. Similarly, diuron is known to block the flow of electrons through photosystem II and thus to stimulate oxidative activity and oxidative stress. However, the disturbances within the redox system were linked to an increased activity of the antioxidant defense mechanisms rather than to increased ROS production [18]. The low effect observed in this study is consistent with the lack of effect of diuron on CAT activity, an important enzyme in cellular metabolism of peroxides in the green microalga Scenedesmus obliquus [42].

Interferences Between C11-BODIPY581/591 Measurements and Micropollutant Induced Shift in Autofluorescence

C. reinhardtii also exhibits autofluorescence observable in the green and orange channels. Orange fluorescence is much less intense in comparison with the chlorophyll autofluorescence observed in the red channel. Green and orange autofluorescence intensity, as well as the red one, can be subjected to variations correlated to changes in algal physiological status and to pollutant triggered stress. To verify if the micropollutant induces a shift in algal green autofluorescence and if that could interfere with measurement of C11-BODIPY581/591 stained cells, FCM analysis of cell autofluorescence of unstained C. reinhardtii but exposed to the micropollutants was also performed. This analysis revealed a shift toward lower values of green autofluorescence in cells exposed for 24 or 48 h to 6 × 10–6 M Cu2+ (Fig. 5). Because of this shift and to correctly discriminate between the effect of micropollutant on the algal autofluorescence detectable in the green channel and fluorescence of the oxidized C11-BODIPY581/591, the gate was transposed to lower green values. Changes in green autofluorescence can be associated with changes in the physiological state of the algal cell [43] and the redox status of pyridine nucleotides in C. reinhardtii [44]. Therefore, algal autofluorescence has to be taken into account when the fluorescent dye is applied in order to avoid over- or underestimation of the dye response.

image

Figure 5. Cell count versus green fluorescence plot. (A) Unstained algae exposed to 6 × 10–6 M Cu2+ for 48 h (pink) and unexposed control (orange) and (B) algae exposed to Cu2+ for 48 h unstained (pink) and stained with C-11-BODIPY581/591 (green). Cells were stained with 2.5 μM C11-BODIPY581/591 for 30 min. [Color figure can be viewed in the online issue which is available at wileyonlinelibrary.com.]

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Conclusion

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

This work fills the existing methodological gap concerning the use of the fluorescent dye C11-BODIPY581/591 in algal cell toxicology and highlighted the suitability of this dye to probe micropollutant induced lipid oxidation in a wide range of ecotoxicological tests involving green algal cells. C11-BODIPY581/591 is a promising dye for algal investigation, because its fluorescence does not interfere with the chlorophyll autofluorescence, it is not toxic to algae and relatively insensitive to variations in dye staining times and algal cell densities. Negligible formation of algal debris and dye precipitation, as well as the stability and reproducibility of the fluorescence measurements make it a reliable stain. Additionally, its specificity of reaction with oxyl-radicals and its stability inside the algal cell guarantee its applicability in a wide range of tests, including studies of lipid oxidation evolution in short term exposures. By coupling C11-BODIPY581/591 staining with flow cytometer analyses, it is possible to obtain, in a single measurement, multiparameter information on cell properties and lipid oxidation of cell membranes.

ACKNOWLEDGMENTS

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Warm thanks are extended to S. Le Faucheur for the help with micropollutant choice and to N. von Moos for helpful discussions on micropollutant-induced oxidative stress.

Literature Cited

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Supporting Information

  1. Top of page
  2. Abstract
  3. Material and Methods
  4. Results and Discussion
  5. Conclusion
  6. ACKNOWLEDGMENTS
  7. Literature Cited
  8. Supporting Information

Additional Supporting Information may be found in the online version of this article.

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