Article pubs.acs.org/est
Cite This: Environ. Sci. Technol. XXXX, XXX, XXX−XXX
N‑Halamine Derivatized Nanoparticles with Selective Cyanocidal Activity: Potential for Targeted Elimination of Harmful Cyanobacterial Blooms Giji Sadhasivam,† Chen Gelber,‡ Varda Zakin,† Shlomo Margel,‡ and Orr H. Shapiro*,† †
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Department of Food Science, Institute of Postharvest and Food Sciences, Agricultural Research Organization, The Volcani Center, Rishon LeZion 7528809, Israel ‡ Department of Chemistry, The Institute of Nanotechnology and Advanced Materials, Bar-Ilan University, Ramat-Gan 52900, Israel S Supporting Information *
ABSTRACT: Harmful cyanobacterial blooms (HCBs) are becoming a major challenge for the management of both natural and man-made freshwater lakes and reservoirs. Phytoplankton communities are an essential component of aquatic ecosystems, providing the basis for natural food webs as well as important environmental services. HCBs, driven by a combination of environmental pollution and rising global temperatures, destabilize phytoplankton communities with major impacts on aquatic ecology and trophic interactions. Application of currently available algaecides generally results in unselective elimination of phytoplankton species, disrupting water ecology and environmental services provided by beneficial algae. There is thus a need for selective cyanocidal compounds that can eliminate cyanobacteria while preserving algal members of the phytoplankton community. Here, we demonstrate the efficacy of N-halamine derivatized nanoparticles (Cl NPs) in selectively eliminating cyanobacteria, including the universal bloom-forming species Microcystis aeruginosa, while having minimal effect on co-occurring algal species. We further support these results with the use a simple microfluidic platform in combination with advanced live-imaging microscopy to study the effects of Cl NPs on both laboratory cultures and natural populations of cyanobacteria and algae at single cell resolutions. We note that the Cl NPs used in this work were made of polymethacrylamide, a nonbiodegradable polymer that may be unsuitable for use as a cyanocide in open aquatic environments. Nevertheless, the demonstrated selective action of these Cl NPs suggests a potential for developing alternative, biodegradable carriers with similar properties as future cyanocidal agents that will enable selective elimination of HCBs.
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hepatotoxins microcystin, nodularin, and cylindrospermopsin.8 Microcystins (MCs) in particular are documented in most of the toxicity incidents associated with HCBs. Acute exposure to these compounds via contaminated water or food may lead to severe liver damage,9,10 while chronic exposure was shown to be carcinogenic, with increased risk of several types of cancer, particularly of the liver or the intestines.11,12 Recognition of the risks associated with MCs led to recommendations of the World Health Organisation (WHO) for a tolerable daily intake (TDI) of no more than 0.04 μg MC per kg body weight per day. This recommendation, which translates to 1 μg/L in drinking water,13 has been adopted by many water authorities worldwide. As microcystine concentrations in HCBs affected water bodies are frequently found to exceed the WHO advisory
INTRODUCTION Harmful cyanobacterial blooms (HCBs) impact lakes, estuaries, and freshwater aquaculture systems worldwide. The duration, severity, and spread of HCBs have markedly increased over the past decades and are likely to increase further in the future, due to the combined effects of anthropogenic-induced eutrophication processes and rising environmental temperatures.1−3 HCBs directly impact the ecology of affected water bodies by altering local food webs, as well as by the release of various secondary metabolites including a diverse suite of toxic compounds collectively termed cyanotoxins.4−7 HCBs are also associated with the release of malodorous compounds, particularly geosmin and 2methylisoborneol, which are considered as major nuisances due to the unpleasant earthy and musty off-flavors that they cause in affected drinking water and aquaculture-grown fish.2 Cyanotoxins are broadly classified as hepatotoxins, neurotoxins, or dermatotoxins, according to their mode of action. Of these, the most commonly observed and monitored are the © XXXX American Chemical Society
Received: Revised: Accepted: Published: A
March 4, 2019 June 8, 2019 June 19, 2019 June 19, 2019 DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Environmental Science & Technology limits,14−16 cyanobacterial blooms have since become a major challenge for water quality management. In recent years, increasing attention has been paid to problems posed by HCBs in fresh water aquaculture (FWA) ponds. FWA is the fastest growing food production sector, with a growth rate exceeding 8% per year, providing over 7% of global consumption of animal protein.17,18 The majority of FWA is based on extensive open ponds, also termed green water operations, where the growth of a diverse community of phytoplankton gives the water a characteristic greenish-brown taint. Phytoplankton growth is generally considered as positive for cultured fish, providing a natural food source, improving water oxygenation, and reducing the accumulation of ammonia which may become toxic for fish at high concentrations.19,20 However, the high nutrient loads in FWA ponds are highly conducive to the development of HCBs, particularly under elevated temperature conditions. While toxin levels rarely result in fish mortality,21 toxin accumulation in fish edible tissues may rapidly reach concentrations that make them unfit for human consumption.22−24 Thus, the monitoring and control of developing HCBs in FWA aquaculture is likely to become a basic requirement for pond management. Multiple treatment strategies, including physical methods and chemical algaecides, are used to control HCBs and their metabolites. While chlorination is generally considered as the fastest and most cost-effective algaecide for bloom management, chlorine toxicity toward fish, and other aquatic organisms, limits its application in aquaculture operations and many other aquatic habitats.25 Herbicidal compounds such as Diuron or Endothall, previously considered as safe alternatives that act specifically on photosynthetic organisms, were recently shown to have nonspecific activity toward nonphotosynthetic biota, as well as potential toxicity of degradation products, limiting their use in drinking water or aquaculture environments.26,27 Copper-based compounds, which are the most commonly used algaecides, are also problematic as ongoing treatment may result in the accumulation of toxic copper ions in the sediments. The nonspecific activity of these algaecides, targeting both algae and cyanobacteria, presents further challenges by disrupting essential ecological services provided by phytoplankton communities in the treated reservoir.28−30 Hence, there is a pressing need for developing cyanocidal agents that enable targeted elimination of cyanobacteria while minimizing the impact on other phytoplankton species and on the aquatic ecosystem as a whole. Recent advances in nanotechnology led to the development of a range of nanomaterials with biocidal properties. The large surface area of nanoparticles (NPs) enhances their interaction with targeted microbes. On the basis of this unique property, NPs of silver, cerium oxide, and zerovalent iron were previously demonstrated as potent algaecides.31−33 However, shortcomings associated with selectivity and large-scale applicability, as well as safety concerns, hinder the endorsement of such nanocides for environmental applications.27 Recently, N-halamine-based antimicrobial agents have attracted a great deal of interest due to their broad spectrum of activity, nontoxicity to humans, good stability, and low cost.34 N-halamine inactivates microorganisms through the direct transfer of positive halogen (Cl+) to the cell membrane receptor leading to cell death.35 An emerging class of nanomaterials that combine the biocidal properties of Nhalamine with the unique physical properties of NPs is
currently being explored as stable, environmentally safe antimicrobial agents for various applications.36−38 Here, we demonstrate the specific activity of N-halamine derivatized polymethacrylamide NPs (Cl NPs) against cyanobacterial cells. Cl NPs were previously shown to have bactericidal activity mediated by the release of oxidizing chlorine ions upon contact with microbial cells, combined with high chemical stability under high organic loads.39,40 The potential of Cl NPs as cyanocides was tested against Microcystis aeruginosa, a universal bloom-forming cyanobacterial species, as well as a suite of additional cyanobacterial and algal strains. Activity was further validated against naturally occurring colonies from HCBs in fish ponds and other freshwater reservoirs. A newly presented approach combining simple microfluidics and live imaging microscopy allowed direct visualization of Cl NPs effects on cyanobacterial cells, using native photosynthetic pigments as natural fluorescent reporters. We propose that future nanomaterials, developed on the basis of the unique properties of the Cl NPs used here, will provide environmentally friendly cyanocides that facilitate targeted elimination of blooming cyanobacteria while minimizing the effect on cohabiting phytoplankton species.
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MATERIALS AND METHODS Preparation of the Chlorinated Nanoparticles. Nhalamine derivatized cross-linked polymethacrylamide NPs (P (MAA−MBAA)) were prepared as described previously by surfactant-free dispersion polymerization mechanism.39 Briefly, P (MAA−MBAA) NPs were prepared by dissolution of 4.4 g of methacrylamide (MAA), 3.6 g of N,N′-methylenebis(acrylamide) (MBAA) (2% w/v total monomers), and 240 mg of potassium persulfate (PPS) in 400 mL of distilled water; the mixture was stirred (200 rpm) at 100 °C for 1 h, yielding particles with an average hydrodynamic diameter of 27 ± 3 nm. MAA and MBAA residues were subsequently removed from the NPs aqueous dispersion by extensive dialysis against water. The chlorinated NPs were prepared by adding sodium hypochlorite aqueous solution (5 mL, 4% w/v) to an aqueous dispersion of the P(MAA−MBAA) NPs (5 mL, 15 mg/mL), and the mixture was shaken at room temperature for 1 h. Excess sodium hypochlorite was removed from the P(MAA− MBAA)-chlorinated NPs dispersion by extensive dialysis against water. The bound-Cl content of the P (MAA− MBAA) chlorinated NPs (Cl NPs) was determined by adding sodium iodide to the Cl NPs dispersion and measuring the resulting color by spectrophotometry at 292 and 350 nm according to the literature.41,42 Phytoplankton Cultures and Growth Conditions. M. aeruginosa PCC 7806 was received from the laboratory of Prof. Aharon Kaplan at the Hebrew University of Jerusalem, Israel, and used as the reference strain for the bulk of our experiments. Additional cyanobacterial strains (Pseudanabaena sp., Limnothrix sp.), diatom (Nitzschia sp.), and algae (Desmodesmus sp., Monoraphidium sp.) were isolated from water samples of several aquaculture ponds and freshwater reservoirs in northern and central Israel (Table S1). Isolates were obtained through successive streaking onto BG 11 media solidified with 0.8% bacteriological agar.43 For isolation of diatom (Nitzschia sp.), BG11 was supplemented with sodium metasilicate (Na2SiO3) to a final concentration of 0.056−0.28 g/L.44 Cultures were maintained at room temperature in BG 11 liquid medium (Sigma-Aldrich) (with added silica for diatoms) under white/blue fluorescent illumination (30 μmol B
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Environmental Science & Technology photons · μm−2 · s−1) with an 18:6 h light: dark cycle. Monospecific cultures of microalgae were identified by morphological appearance under light microscopy as well as by partial sequencing of rRNA. The universal forward primer 27F45 and cyanobacteria specific reverse primer CYA784R46 was used for 16S rRNA based identification in cyanobacteria, and microalgae specific P73F and P47R for 18S rRNA identification in algae and diatom.47 Natural HCB Samples. Samples were obtained from freshwater reservoirs with an active HCB bloom resembling Microcystis sp. HCB1 was collected from an aquaculture pond in the north of Israel and kept in the laboratory for 24 h prior to experiment; HCB2 & HCB3 were collected from a freshwater reservoir in the center of Israel and used for experiment within 2 h of collection. Effect of Cl NPs on Microcystis aeruginosa. Cl NPs was tested using the toxic cyanobacterium strain M. aeruginosa PCC 7806 as a model organism. A suspension of M. aeruginosa (5 × 105 cells/ml) in 20 mL BG11, held in 50 mL sterile culture flasks, was challenged with Cl NPs at different chlorine concentrations (0.1 mM, 0.5 mM, and 1 mM). Cl NPs from a stock solution of known concentration (28 mM, determined as described above) were diluted (0.1 mM (71.4 μL/20 mL), 0.5 mM (357 μL/20 mL), and 1 mM (714 μL/20 mL)) to desired concentration in M. aeruginosa culture in BG11. Cells treated with an equal volume of dw/non chlorinated NPs (stock of 14 mg/mL non chlorinated NPs equivalent to 28 mM Cl NPs) served as control. The experiment was performed in triplicate with 18:6 h light: dark under white/blue fluorescent illumination (30 μmol photons · μm−2 · s−1) at room temperature for 10 days. Changes in autofluorescence of photosynthetic pigments (chlorophyll and phycocyanin) were measured by daily sampling (200 μL) from each flask. Samples were transferred into a 96 well microtiter plate and autofluorescence of the native pigments chlorophyll (Ex: 485 nm/Em: 680 nm) and phycocyanin (Ex: 590 nm/Em: 650 nm)48 was measured using a microplate reader (Biotek’s synergy Neo2 multimode). All fluorescence measurements were done in triplicates and expressed as arbitrary fluorescence units. Comparison between Sodium Hypochlorite and Cl NPs. The effects of sodium hypochlorite and Cl NPs on cyanobacteria (M. aeruginosa PCC 7806) and algae (Desmodesmus sp. isolated from a fish pond in northern Israel) were tested by incubating cultures of equal cell concentrations (4 × 106 cell/ml) with different chlorine concentrations (0.01, 0.1, and 1 mM) derived from either 3% bleach (commercial) or Cl NPs (as described above). The assay was performed in a 96 well microtiter plate, by adding to each well 100 μL of cell suspension (8 × 106 cells/ml) in BG11, and an equal volume of either bleach or Cl NPs diluted in dw to twice the final concentration. Each combination was tested in triplicates. Cultures treated with 100 μL of dw served as control. To monitor the immediate effects of treatments, autofluorescence of the photosynthetic pigments (chlorophyll and phycocyanin, as described above) were measured immediately following treatment and then at 5 min intervals over the next 80 min. Microtiter plates were then incubated under conditions as described above, and readings were obtained at daily intervals over the next 10 days. ζ Potential Measurement. The surface charge of algae, cyanobacteria, and Cl NPs were analyzed using Zetasizer Nano ZS (Malvern instruments, UK) with a light source of He−Ne
laser 633 nm. Cells of algae (Desmodesmus sp) and cyanobacteria (M. aeruginosa) were harvested by centrifugation at 3354 X g for 10 min at 20 °C. The cell pellets were washed twice in fresh BG11 medium by repeating the centrifugation step. The final cell pellet was resuspended in BG11 medium and used for analysis. Algae and cyanobacteria cells suspension treated with 1 mM of Cl NPs (Test) or BG11 (Control) were examined for ζ potential at 2 and 24 h following treatment. Measurements were performed in triplicates for each sample (3 scans, 100 runs each), with an initial equilibration time of 5 min at 25 °C using the Malvern’s DTS software. Mean ζ potential values for each sample were obtained. Microfluidic Chamber Construction. Elongated hexagonal chambers (10 mm × 4 mm × 0.15 mm; Chamber volume 6 μL) were etched into a silicone elastomer (polydimethylsiloxane (PDMS)) (Sylgard 184, Dow Corning) using soft lithography.49,50 Inlet holes were punched at both ends of each well using a 1 mm biopsy punch (AcuDerm, FL, USA). Channels were placed on the clean surface of a new glass microscope slide and bonded by 30 s activation of both surfaces using a corona discharge torch (Elecrotechnique products, USA) followed by heat treatment (15 min at 100 °C). Microfluidic chambers were placed in a vacuum chamber prior to experiment to minimize the formation of bubbles that may interfere with microscopy. Live Imaging Microscopy Experiments. To visualize the effect of Cl NPs on different strains or on natural HCBs, a suspension of the desired strain or HCB sample was diluted with an equal volume of either Cl NPs (final concentration of 10 mM chlorine) or distilled water for control wells. An aliquot from each sample was loaded into a microfluidic chamber using a 10 μL pipette, and the chamber covered with a thin film of PDMS to minimize evaporation through the inlet holes. Microscopic imaging was performed using a NIKON eclipse Ti microscope (Nikon, Japan) equipped with a ProScan motorized XY stage and an HF110A system (enabling rapid switching of emission filters)(Prior Scientific, MA, USA), and a temperature-controlled stage incubator (LAUDA ECO RE 415, Korea). Bright field illumination was provided by a cool LED pE-100A (Cool LED, UK). Excitation light for epifluorescence microscopy was provided by a Spectra X light engine (Lumencor, USA). Imaging was performed using a long working distance 40x objective (NA 0.6) (Nikon, Japan). Images were captured at 30 min intervals using an ANDOR zyla 5.5 MP sCMOS camera (Oxford Instruments, UK). For each frame, bright field (gray), phycocyanin (Ex: 555 nm/Em: 632 nm; pink) and chlorophyll (Ex: 440 nm/Em: 697 nm; red) were captured separately and overlaid to produce the final image. Images were processed using the NIS elements AR 4.6 (64 bit) software package (Nikon, Japan).
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RESULTS Effect of Chlorinated Nanoparticles on Microcystis aeruginosa. Chlorinated nanoparticles (Cl NPs) activity against M. aeruginosa showed a dose-dependent kinetics. For all treated samples, an increase in phycocyanin fluorescence was observed, followed by rapid decline in autofluorescence of both chlorophyll and phycocyanin (Figure 1). No such decline was observed for control samples. The timing of the observed response varied between 24 h for 1 mM to 3−4 days for 0.1 mM chlorine. Nonchlorinated polymethacrylamide (MAA− MBAA) NPs at a concentration of 7 mg/mL (over 10 times C
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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returned a ζ potential value (ZPV) of approximately −28 mV, compared to a ZPV of −17 mV for Desmodesmus sp. At 2 h following treatment with Cl NPs, ZPV of M. aeruginosa shifted to −23 mV, and a further shift towards −21 mV was measured at 24 h following treatment. No significant difference was found between treated and nontreated Desmodesmus cells. Microscopic Observation and Analysis of Cl NPs Activity. Kinetics of Cl NPs effect on M. aeruginosa was visualized using live-imaging microscopy. Using the native fluorescence of chlorophyll and phycocyanin we were able to track changes in the viability of individual cells in a culture maintained within a simple microfluidic device over a period of 24−48 h. In M. aeruginosa cells treated with Cl NPs (10 mM chlorine), a slight increase in phycocyanin fluorescence was observed within 4−5 h of treatment, followed by prolonged signal decay. Loss of autofluorescence was frequently preceded by sudden deformation in cell structure (e.g., cell 2 in Figure 3; see also Supporting Information (SI) video S1), confirming that loss of fluorescence following Cl NP treatment is indeed indicative of cell death. The selective activity of Cl NPs against additional cyanobacterial and algal species was similarly visualized using live-imaging microscopy. No changes in chlorophyll autofluorescence were observed in the eukaryotic algae Monoraphidium sp. and Desmodesmus sp. (Figure 4A,B; SI video S2 (Desmodesmus sp.)). Surprisingly, complete loss of chlorophyll fluorescence was observed in a diatom strain identified as Nitzschia sp. within 2 h of exposure to Cl NPs (Figure 4C; SI video S3). A similar response was observed in unidentified diatom cells found in natural samples (results not shown), suggesting high sensitivity of diatoms to Cl NPs. Additional cyanobacterial strains identified as Pseudanabaena sp. and Limnothrix sp. showed a similar response to that observed with M. aeruginosa following Cl NPs treatment (Figure 4D,E; SI video S4 (Pseudanabaena sp.)). Differences in the response time may represent differences in sensitivity among the tested strains. The selectivity of Cl NPs was further demonstrated using the microfluidic approach by treatment of a mixture of two algal species (Monoraphidium sp. and Desmodesmus sp.) and two cyanobacteria strains (M. aeruginosa and Pseudanabaena sp.) (Figure 4F; SI video S5). Activity of Cl NPs was further tested against naturally occurring HCBs (termed here (HCB1-HCB3) collected from aquaculture ponds and other freshwater reservoirs in Israel. All tested colonies were sensitive to Cl NPs. However, some
the concentration used for the 1 mM treatment) had no effect on M. aeruginosa culture over a period of 7 days (Figure S1). Selective Activity of Cl NPs against Cyanobacteria. The selectivity of Cl NPs against cyanobacteria was tested by comparing activity of bleach and Cl NPs against the cyanobacterium M. aeruginosa PCC7806 and an algal strain identified as Desmodesmus sp. Addition of 0.1 or 1 mM chlorine as resulted in the rapid loss of autofluorescence of in both Desmodesmus sp. and M. aeruginosa within 10 min of treatment (Figure 2A−C, insets). Some level of selectivity was found following the addition of 0.01 mM chlorine as bleach, where no effect was observed for Desmodesmus cells, while the M. aeruginosa culture collapsed at 3−4 days following treatment (Figure 2A−C). Unlike bleach, a clear selectivity was found for treatment with Cl NPs at similar chlorine concentrations. Although some inhibition of Desmodesmus growth was observed following treatment with 0.1 and 1 mM chlorine, none of the treatments resulted in the collapse of Desmodesmus cultures (Figure 2D). Contrastingly, M. aeruginosa again responded in a dosedependent manner, with the culture collapsing within 1−4 days of treatment (Figure 2E,F). Similar selectivity of Cl NPs activity was demonstrated for other algal and cyanobacterial species (Supporting Information, Table S2). ζ Potential Measurement. ζ potential measurements for Cl NPs as well as treated and nontreated M. aeruginosa and Desmodesmus cells are given in Table 1. Cl NPs were found to Table 1. Changes in ζ potential values of M. aeruginosa and Desmodesmus sp cells following Cl NPs treatment Sample details 1 mM Cl NPs suspended in BG11 M. aeruginosa (Control) M. aeruginosa (Treated) Desmodesmus sp. (Control) Desmodesmus sp. (Treated)
ζ potential (mV) (2 h incubation)
ζ potential (mV) (24 h incubation)
−12.1 ± 0.9
−12.1 ± 1.7
−28.0 ± 2.2 −23.2 ± 2.3 −18.3 ± 1.3
−27.5 ± 3.1 −20.5 ± 1.7 −16.9 ± 1.2
−19.0 ± 1.8
−18.2 ± 2.0
be negatively charged (approximately −12 mV), similar to previous results.40 The surface charges of both algae and cyanobacteria were also found to be negative under the conditions tested (pH 6−7). Untreated M. aeruginosa cells
Figure 1. Dose-dependent effect of Cl NPs on M. aeruginosa PCC7806. Changes in autofluorescence of chlorophyll (A) and phycocyanin (B) following treatment with 0.1, 0.5, and 1 mM of Cl NPs. Decrease in the autofluorescence of chlorophyll or the observed rise and subsequent decrease for phycocyanin marks culture collapse. All measurements were normalized to time ‘0’ for ease of interpretation. D
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Figure 2. Selective cyanocidal effect of Cl NPs. Effects of Cl NPs and bleach at three different concentrations tested against algae (Desmodesmus sp.) and cyanobacteria (M. aeruginosa). (A−C) Algae and cyanobacteria responded in a similar manner to bleach concentrations of 0.1 and 1 mM. Inset shows measurements during initial 80 min following treatment. (D−F) Differential action of Cl NPs against Desmodesmus (part D) vs Microcystis (parts E and F).
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differences were observed between the kinetics of the different colonies (Figure 5A−C). HCB1 showed rapid inflation of the colony, possibly through oxidation of extracellular polymeric substances (EPS) holding the cells together. This was followed by simultaneous loss of phycocyanin fluorescence of all cells in the colony, with near complete loss of fluorescence 15 h following treatment (Figure 5A,B; SI video S6 (HCB1)). Colonies from HCB2 appeared to be surrounded by a thick mucilaginous sheath. Here, colony integrity was maintained and individual cells responded at different times following Cl NPs treatment, with each cell showing a sharp increase in phycocyanin autofluorescence followed by loss of the fluorescence signal (Figure 5C,D; SI video S6 (HCB2)). Interestingly, this response was observed over a period of 40 h, demonstrating the prolonged activity of Cl NPs following adsorption to a cyanobacterial colony. A different kinetic response was observed in colonies from a third bloom (HCB3). Here, phycocyanin fluorescence remained stable for the first 10 h following Cl NPs treatment. A rise in fluorescence of all cells in the colony was observed between 10 and 13 h following treatment, and this was followed by a gradual loss of signal over the next 15 h (Figure 5E,F; SI video S6 (HCB3)). No change was observed in colony structure following treatment. Autofluorescence of control, untreated colonies from the same blooms, maintained under similar conditions but without the addition of Cl NPs, appeared stable over the same time period (data not shown).
DISCUSSION
The control of bloom-forming cyanobacteria and the elimination of the health risks associated with HCBs are becoming major challenges on a global scale. Long-term solutions for these problems will necessary involve better environmental management and reduced pollution and nutrient loads in affected water bodies. However, currently available solutions primarily rely on the application of algaecides as a means to curb developing blooms. Commercially available algaecides utilize either general herbicidal chemicals (e.g., diuron), oxidizing agents, (e.g., chlorine or hydrogen peroxide), or copper-based compounds.27 While such measures are generally effective, there are many reports of algal and cyanobacterial communities developing resistance to herbicides and copper-based algaecides following repeated use.51,52 One problem with currently available algaecides is their nonspecific action targeting both eukaryotic algae and cyanobacteria and often affecting additional components of the ecosystem.51,27 In many environments, including freshwater aquaculture operations, such a general biocidal effect is undesirable. Phytoplankton communities play a central role in aquatic ecology, e.g., in water oxygenation, nutrient recycling, and as the basis of multiple food webs. Similarly, extensive FWA operations rely on these activities to maintain pond water quality and ensure the wellbeing of cultured fish.19,20 There is thus a real need to develop novel compounds or formulations with selective cyanocidal activity that will allow treatment of E
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Figure 3. Cl NPs effect on single cells of M. aeruginosa. (A) Representative frames from a 24 h live imaging microscopy sequence. Loss of phycocyanin fluorescence (magenta) is observed in all cells in the frame. Cell 2 shows cell deformation (6:31 h) preceding the loss of phycocyanin fluorescence. (B) No changes in cell shape or phycocyanin fluorescence were observed in control wells. (C) Changes in phycocyanin fluorescence (normalized) measured for single cells (numbered 1−5 in A). (D) Averaged changes in fluorescence for control and Cl NPs treated M. aeruginosa cells (data from average of 10 randomly selected cells for each treated and the control). Error bars indicate standard error.
compounds, which are widely used as disinfectants and algaecides, are generally considered nonselective,55 the observed selectivity in our experiments is likely to be a property of the carrier particles rather than the functional groups. While differential susceptibilities of cyanobacterial and algal cells to oxidative stress, known from previous studies56,57 and seen also in our bleach treatment (Figure 2A−C), may account for some of the observed difference, it does not explain the much higher activity of Cl NPs toward cyanobacterial cells or the unique kinetic response of the treated cells (Figure 2D−F). Unlike the addition of bleach, where concentrations exceeding 0.1 mM chlorine induced the rapid loss of autofluorescence for both cyanobacterial and algal cells, addition of Cl NPs had no immediate effect on either culture but induced the collapse of cyanobacterial cultures within 24− 72 h depending on concentration. Interestingly, a similar effect on M. aeruginosa was observed following the addition of 0.01 mM chlorine as bleach, a concentration that was too low to affect the Desmodesmus culture. Importantly, a high level of selectivity was observed also when algae and cyanobacteria were treated in a mixed culture with a high Cl NPs concentration of 10 mM chlorine (i.e., 100 times higher than the concentration needed to bleach algal cultures (Figure 2A)) (Figure 4F; SI video S5). This result indicates that Cl+ ions, upon dissociation from Cl NPs, are immediately consumed, as diffusion of even 1% of free Cl+ ions to the surrounding environment should also have affected neighboring algal cells. From this result, we may conclude that the interaction of Cl NPs with cyanobacterial cells is likely to occur at the vicinity of the cell surface, as demonstrated previously for bacterial cells,40 facilitating the immediate
HCBs with minimal disturbance to algae and other aquatic species.27 The results presented here are the first demonstration of the use of chlorinated nanoparticles as cyanocidal agents. We demonstrate the efficacy and selective activity of Cl NPs against several cyanobacterial and algal strains, including a model laboratory culture (M. aeruginosa PCC 7806) as well as less domesticated strains recently isolated from freshwater aquaculture ponds. Moreover, we find similar activity against cyanobacterial microcolonies that were collected from natural blooms and maintained in the water from which they were collected, demonstrating the potential of Cl NPs to be used under conditions resembling those found in natural habitats. It is important to note that the particles used in this study are based on polymethacrylamide. This polymer is considered nontoxic to plants and animals and is used in bulk quantities as a flocculation agent in water and wastewater treatment, as a soil stabilizer in agriculture, and as a viscosity modifier in the oil industry.53 Nevertheless, polyacrylamides are considered nonreadily biodegradable54 and, under UV exposure, they may be degraded into the toxic and potentially carcinogenic acrylamide.53 Its introduction in the form of nanoparticles into surface waters, particularly into food production environments such as aquaculture ponds, should thus be treated with some caution. A central question arising from our results is that of the mechanism underlying the observed selectivity. Cl NPs kill cells by releasing Cl+ ions, similar to sodium hypochlorite or bleach.39,40 The activity potential of Cl NPs should thus depend primarily on Cl+ density, i.e., the concentration of Nhalamine moieties per surface area. 40 As N-halamine F
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Figure 4. Microscopy based tracking of Cl NPs activity against algae, diatom, and cyanobacteria. Live imaging microscopy and subsequent image analysis was used to test the effect of Cl NPs against isolated strains (A−E) and artificial communities (F). The artificial community shown in part F includes two algal species (Monoraphidium sp., Desmodesmus sp.) and two cyanobacterial species (M. aeruginosa and Pseudanabaena sp.) (data presented is the normalized average of 10 cells/species; single cell tracking was performed using either phycocyanin for cyanobacteria or chlorophyll for algae).
charged Cl NPs, resulting in the observed reduction in ζpotential value. Such a preferential attachment of Cl NPs to cyanobacterial cells may be mediated through differences in the cell wall composition,65,66 e.g., due to the presence of carboxyl groups mediating the adsorption of cations on the surface of M. aeruginosa65 or through some other mechanism to be explored in future experiments. In our bulk measurements of cyanobacterial cells treated with Cl NPs, we consistently observed an increase in phycocyanin autofluorescence followed by decay of fluorescence signal from both phycocyanin and chlorophyll. As noted above, the timing of this response depended on Cl NPs concentration. A similar response was observed for M. aeruginosa immediately following treatment with 0.1 mM chlorine as bleach (Figure 2C). This effect was described previously following chlorination67 or sonication67,68 of cyanobacterial cells and is attributed to the dissociation of phycocyanin from photosystem I/II, resulting in an increase in the free phycocyanin and its specific fluorescence.67 This characteristic response has recently been proposed as the basis for a monitoring system for the state of cyanobacterial blooms in natural environments.67 During the present study, we developed a novel screening approach combining live imaging fluorescent microscopy with a simple microfluidic platform. Similar to our bulk experiments, we utilize the autofluorescence of photosynthetic pigments, and particularly the observed shift in phycocyanin fluorescence following stress, to study the dynamic effect of Cl NPs on
consumption of free Cl+ ions. Furthermore, this result suggests that Cl NPs act by releasing their Cl+ cargo over an extended time period, as rapid release should result in more free ions “escaping” to the surrounding environment. Such a slow release mechanism could induce a continuous state of sublethal oxidative stress in affected cells ultimately resulting in cell demise, possibly similar to a previously demonstrated mechanism in the green algae Chlamydomonas reinhardtii, where two consecutive events of sublethal oxidative stress induced culture collapse via programmed cell death.58,59 We hypothesized that the specific interaction of Cl NPs with cyanobacteria may be mediated by differences in the surface charge of cyanobacterial and algal cells. However, our results do not support such a mode of attraction, as both Cl NPs and cells were found to be negatively charged under the conditions tested (Table 1). Nevertheless, a relatively large positive shift in ζ-potential, from −28 to −23 mV, was observed in M. aeruginosa cells already 1 h following treatment with Cl NPs (Table 1), and a further shift toward −21 mV was observed 24 h later, whereas no such shift was observed in Desmodesmus cells. A similar positive shift is often reported following chlorination of algal and cyanobacterial suspensions60−64 and is attributed to the interaction of positive chlorine ions with the negatively charged surface leading to charge neutralization. However, such a mechanism should lead to a similar effect on both algae and cyanobacteria, which is not observed here. An alternative explanation would be that M. aeruginosa, but not Desmodesmus cells, forms aggregates with the less negatively G
DOI: 10.1021/acs.est.9b01368 Environ. Sci. Technol. XXXX, XXX, XXX−XXX
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Figure 5. Effect of Cl NPs on naturally occurring HCB colonies. Changes in phycocyanin fluorescence of single cells in natural HCB colonies treated with Cl NPs were tracked using live imaging microscopy. (A, C, E) Changes in the autofluorescence of individual cells within treated colonies. (B,D,E) Snapshots from the image sequneces used for measuring data in A,C and E, respectively, showing differences in colony morphology and temporal response to Cl NPs treatment.
results with our Nitzchia isolate (data not shown), suggesting the observed sensitivity of Nitzchia to represent a more general trait of at least some diatom lineages. Possible mechanisms behind this increased sensitivity may be different cell-surface properties of diatoms promoting adhesion of Cl NPs, increased sensitivity to oxidative stress compared to other microalgae, or the relatively large surface area to biomass ratio (due to the presence of a large vacuole) which would result in a proportional increase in oxidative stress due to interactions with Cl NPs. The unique effect of Cl NPs on diatoms further highlights the difference between them and other algaecidal agents. While the selectivity between cyanobacteria and algae does not hold for diatoms, the loss of diatoms from the phytoplankton community may be considered a relatively low price to pay for the elimination of HCB’s. Indeed, diatoms may themselves form nuisance blooms69−71 and biofouling in both freshwater and marine environments, and their observed sensitivity to Cl NPs may thus be considered, at least under some scenarios, advantageous. Certainly, this aspect of Cl NPs activity should be explored in future experiments. An interesting outcome from our microfluidic experiments was that the kinetics of phycocyanin autofluorescence observed in the bulk experiments, i.e., the relatively sharp increase and subsequent prolonged decay in signal intensity, is largely reproduced at the level of individual cells (Figures 3 and 4). This suggests that the bulk observation does not necessarily
cyanobacteria and alga at single cell resolution. The small and confined volume of the chambers used in our system minimizes convective mixing of the sample, allowing us to use time-lapse microscopy to track the fate of individual cells throughout each experiment. The small chamber volume, ca. 10 μL, further allowed us to run multiple experiments using a high concentration of Cl NPs, expediting results while saving the need to use large quantities of Cl NPs. Results from this system further confirmed the selective activity of Cl NPs against cyanobacteria. A near complete collapse of M. aeruginosa cultures was observed within 10−20 h of addition of Cl NPs and a similar result was observed in several additional cyanobacterial strains. In contrast, and despite the high (10 mM) chlorine concentration used in these experiments, no effect was observed in four eukaryotic algae tested following a 48 h exposure to Cl NPs. A notable exception to the observed selectivity was the high sensitivity of an isolated diatom, (Nitzchia sp.) to Cl NPs (Figure 4C, SI video 3). On the basis of microscopic analysis, diatoms did not form a major part of any of the natural cyanobacterial blooms we sampled for the current work, and no additional diatom strains were isolated or tested. However, during a couple of microscopy-based experiments we did observe single diatoms attached to microcystis-like colonies. In both these observations, the diatoms’ chlorophyll fluorescence was extinguished early into the experiment, similar to the H
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morphology is perfectly preserved following treatment, similar to HCB2, despite the lack of a visible mucilage sheath. The effect on individual cells appears also to be delayed. However, the peak in auto fluorescence for all cells appears nearly simultaneously, perhaps indicating this delay to be mediated by some inherent resistance to oxidation rather than delayed Cl NPs penetration. Thus, the efficacy of Cl NPs under natural conditions may vary for different bloom types and environmental conditions. To conclude, we demonstrated selective activity of Cl NPs against several cyanobacterial strains, including toxic and potential bloom-forming species, as well as their potential as cyanocidal agents under conditions resembling those found in natural aquaculture ponds or natural reservoirs. While the precise mechanism underlying this specificity remains to be determined, we provide several insights into both the site and mode of interaction of Cl NPs with cyanobacterial cells. We believe that the unique properties of this system justify further exploration and such an effort will enable the informed design of a future generation of environmentally safe nanocyanocides, providing a much needed solution for treating HCBs in aquaculture ponds and other freshwater environments.
represent gradual loss of individual cells but rather a semisynchronized loss of fluorescence of the entire population. Another related observation is that whenever cell deformation was observed (e.g., Figure 3A; SI video S1), it invariably preceded the decay in autofluorescence. For these cells, at least, the observed decay in autofluorescence occurred already following cell death. It is unclear if this is the case also for cells where deformation was not observed, i.e., at what stage of signal decay should cells no longer be considered viable. As far as we can tell, based on our current observations, we did not see a reversal of signal decay in any of treated strains, suggesting that loss of phycocyanin autofluorescence may be an irreversible process resulting from or leading to cell death. Yet another observation from the same set of experiments was that in some of the tested species, the timing of signal decay varied between cells. This variability was relatively low for M. aeruginosa but was more pronounced in filamentous cyanobacterial species (Figure 4D,E). Again, the precise mechanism underlying this variability remains to be explored. We further observed a near complete inhibition of motility in filamentous cyanobacteria following Cl NPs treatment (SI videos 4), indicating either physical arrest of the motility apparatus by attached nanoparticles or a particular sensitivity of that mechanism to oxidative stress. It should be noted that no effect on motility was observed following treatment with nonchlorinated NPs (data not shown). The introduction of naturally occurring cyanobacterial colonies into our microfluidic assay asserted the usefulness of Cl NPs under more natural conditions. It was important to verify that the activity of Cl NPs was not limited to lab-grown cultures in clean media but that they were also effective for cells under the conditions characteristic of naturally occurring HCBs. Under these conditions, many cyanobacteria, including Microcystis, form colonies of different sizes and complexities, facilitating daily vertical migrations in the water column as well as defense against grazing pressure.72−75 Moreover, natural pond water is much more complex than laboratory growth media, containing a variety of microorganisms and organic molecules. Natural blooms may be more resistant to treatment due to the difficulty of NPs to penetrate the complex threedimensional matrix of the colony or loss of oxidative potential due to the interaction of nanoparticles or Cl+ ions with nontarget organisms or with organic matrices. In our experiments, we utilized colonies from natural HCBs, with or without a mucilage sheath which may provide protection from external chemical attack.76 We observed a clear response in all colonies following the application of Cl NPs (10 mM Cl+). Intriguingly, the effects on different colony types had unique patterns. For HCB1, a rapid expansion of the colony was observed immediately following addition of Cl NPs (Figure 5A,B; SI video S6 (HCB1)), likely representing the destruction of extracellular polymers bonding the cells.76,77 The subsequent synchronized loss of auto fluorescence throughout the colony suggests minimal inhibition of penetration of Cl NPs. A very different response was observed for colonies collected from HCB2, which appeared to be enveloped by a mucilage sheath (Figure 5C,D; SI video S6 (HCB2)). Here, the retention of colony morphology suggests that the sheath was not substantially affected by Cl NPs treatment. Cells within the colony are clearly affected but with large temporal variation, suggesting that the sheath acted as a partial barrier for the penetration of Cl NPs. Yet another kinetic is observed for HCB3 (Figure 5E,F; SI video S6 (HCB3)). Here, again colony
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ASSOCIATED CONTENT
* Supporting Information S
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.est.9b01368. List of phytoplankton species and their collection sites, nonchlorinated nanoparticles (PMAA) showing no growth inhibition effect on M. aeruginosa cells, effect of Cl NPs was validated on two other species of algae under microscopy, variation in the cell count of algae and cyanobacteria grown with supplementation of 0.5 mM of Cl NPs in BG11 (PDF) Control and Cl NPs treated M. aeruginosa (MP4) Control and Cl NPs treated Desmodesmus sp. (MP4) Control and Cl NPs treated Nitzschia sp. (MP4) Control and Cl NPs treated Pseudanabaena sp. (MP4) Mixture of algae and cyanobacteria treated with Cl NPs (MP4) Naturally occurring HCBs treated with Cl NPs (MP4)
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Tel: 03-9683422. ORCID
Orr H. Shapiro: 0000-0002-3222-9809 Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The research was funded by grants 20-06-0044 and 20-060065 from the chief scientist of the Ministry of Agriculture and rural development, Israel to O.H.S. and S.M. We thank Prof. Aaron Kaplan, Hebrew University of Jerusalem, Israel, for providing us with the M. aeruginosa strain PCC 7806.
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REFERENCES
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