Detection of cyanobacteria in eutrophic water using a portable

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Detection of cyanobacteria in eutrophic water using a portable electrocoagulator and NanoGene assay Eun-Hee Lee, Beelee Chua, and Ahjeong Son Environ. Sci. Technol., Just Accepted Manuscript • DOI: 10.1021/acs.est.7b05055 • Publication Date (Web): 05 Jan 2018 Downloaded from http://pubs.acs.org on January 10, 2018

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Environmental Science & Technology

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Detection of cyanobacteria in eutrophic water using a

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portable electrocoagulator and NanoGene assay

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Eun-Hee Lee, 2,* Beelee Chua, 1,*Ahjeong Son

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Department of Environmental Science and Engineering, Ewha Womans University, Seoul, Republic of Korea

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School of Electrical Engineering, Korea University, Seoul, Republic of Korea

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*Corresponding Author, Beelee Chua: Present address. 145 Anam-ro, Seongbuk-gu, Korea University, Seoul,

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02841, Republic of Korea; E-mail. [email protected]; Phone. +82 (2) 3290-4639

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*Corresponding Author, Ahjeong Son: Present address. 52 Ewhayeodae-gil, Seodaemun-gu, Ewha Womans

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University, Seoul, 03760, Republic of Korea; E-mail. [email protected]; Phone. +82 (2) 3277-3339; Fax.

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+82 (2) 3277-3275

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ACS Paragon Plus Environment


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ABSTRACT

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We have demonstrated the detection of cyanobacteria in eutrophic water samples using a portable

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electrocoagulator and NanoGene assay. The electrocoagulator is designed to pre-concentrate cyanobacteria from

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water samples prior to analysis via NanoGene assay. Using Microcystis aeruginosa laboratory culture and

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environmental samples (cell densities ranging from 1.7 × 105 to 4.1 × 106 and 6.5 × 103 to 6.6 × 107 cells·mL-1,

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respectively), the electrocoagulator was evaluated and compared with a conventional centrifuge. Varying the

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operation duration from 0 to 300 s with different cell densities was first investigated. Pre-concentration

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efficiencies (obtained via absorbance measurement) and dry cell weight of pre-concentrated cyanobacteria were

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then obtained and compared. For laboratory samples at cell densities from 3.2 × 105 to 4.1 × 106 cells·mL-1, the

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pre-concentration efficiencies of electrocoagulator appeared to be stable at ~60%. At lower cell densities (1.7

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and 2.2 × 105 cells·mL-1), the pre-concentration efficiencies decreased to 33.9 ± 0.2 and 40.4 ± 5.4%,

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respectively. For environmental samples at cell densities of 2.7 × 105 and 6.6 × 107 cells·mL-1, the

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electrocoagulator maintained its pre-concentration efficiency at ~60%. On the other hand, the centrifuge’s pre-

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concentration efficiencies decreased to non-detectable and below 40%, respectively. This shows that the

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electrocoagulator outperformed the centrifuge when using eutrophic water samples. Finally, the compatibility of

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the electrocoagulator with the NanoGene assay was verified via the successful detection of the microcystin

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synthetase D (mcyD) gene in environmental samples. The viability of the electrocoagulator as an in situ

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compatible alternative to the centrifuge is also discussed.

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Keywords: electrocoagulator; centrifuge; pre-concentrate; cyanobacteria; NanoGene assay; Microcystis

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aeruginosa

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INTRODUCTION 2


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In the summer of 2014, a three-day tap water ban was imposed in Toledo, Ohio (population ~ 300,000), due to a

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massive harmful algal bloom on the surface of Lake Erie, USA.1-3 In the following summer, a harmful algal

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bloom along the coast of Washington state resulted in a loss of over 9 million US dollars in razor clam fisheries

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alone.4 According to the National Oceanic and Atmospheric Administration,5 harmful algal blooms have so far

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occurred in every coastal state in the USA. This is because they tend to proliferate in anthropologically altered

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environmental conditions such as eutrophic waters fed by sewage and agricultural runoff.6

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It is well documented that harmful algal blooms can cause acute morbidity and mortality of invertebrate

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and fish species via oxygen depletion (hypoxia and anoxia).7-10 Harmful algal blooms are also responsible for the

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clogging of screens, fouling of weirs, and interference of floc settling in water treatment plants.11-13 They are

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also a source of taste and odor compounds such as geosmin and 2-methylisoborneol and therefore cause

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malodorous and unpalatable drinking water. Most importantly, harmful algal blooms that are primarily

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composed of cyanobacteria and certain common strains of algae are known to produce toxic secondary

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metabolites and endotoxins such as neurotoxins, cytotoxins, dermatotoxins, irritant toxins (lipopolysaccharides),

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and hepatotoxins.14 For example, the genera Microcystis, Anabaena, and Plankthotrix can produce microcystins

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(i.e., hepatotoxins).6 Therefore, chronic and acute exposure to microcystins can result in liver failure and cancer.

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In this regard, in situ cyanobacterial detection, particularly detection of the above-mentioned toxigenic

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genera, would have an obvious benefit. The rapid identification and quantification of specific toxin-producing

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strains during a bloom would be invaluable. It would allow authorities to restrict recreational activities in parks

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and fisheries to protect their stocks and water treatment plants to accommodate a higher than usual toxin load in

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a timely manner. However, existing cyanobacterial detection techniques are not particularly amendable for in

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situ deployment. So far, they have been largely limited to the analysis of field samples using laboratory-based

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equipment (e.g., microscopic inspection, chromatographic and spectroscopic analysis, and molecular biology

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assays).14-18 3



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Furthermore, prior to analysis, the field samples require pre-concentration followed by lysis and

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purification. Currently, pre-concentration can be achieved using a centrifuge or by filtration via a fine membrane.

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Portable centrifuges (such as the Portable Centrifuge Kit by Healthrow Scientific) are electrical power intensive

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(requiring a power adaptor) as well as costly (> 300 USD). Given the small size of cyanobacteria, the use of

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filtration via a fine membrane requires a pump with high operating head and hence is also power intensive and

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costly. Therefore, neither approach is amenable for in situ operation.

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Inspired by the use of electrocoagulation in water treatment and algae harvesting,19-23 we have

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developed a pocket-size portable electrocoagulator suitable for in situ cyanobacterial detection with the

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NanoGene assay. Electrocoagulation is a well-known process that uses electricity to dissolve metal in order to

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supply the ions required for coagulation of colloids including microorganism.24-25 In this study, aluminium was

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used as both anode and cathode in the electrocoagulation process. At the anode surface where oxidation occurs,

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the major products are H+, Al3+, and O2 gas. The corresponding oxidative half reactions are as follows.

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Al → Al3+ + 3e-

(1a)

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2H2O → O2 + 4H+ + 4e-

(1b)

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2OH- → O2 + 2H+ + 4e-

(1c)

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At the cathode surface where the reduction occurs, the major products are OH-, and H2 gas. The corresponding

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reductive half reactions are as follows.

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2H2O + 2e- → H2 + 2OH-

(2a)

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2H+ + 2e- → H2

(2b)

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Aluminium ions (Al3+) are generated at the anode and they form aluminium hydroxide (Al(OH)3). The

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subsequent hydration of Al(OH)3 will further produce a variety of macro ions or gelatinous precipitates, which

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will facilitate the coagulation of colloids such as microorganisms. In addition, gases trapped in coagulated

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microorganisms will cause them to float to the surface.

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The NanoGene assay is a bio-assay based on magnetic beads, dual quantum dots as well as DNA hybridization.

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It is well known for its inhibitor resistance, sensitivity, selectivity, and in-situ compatibility for detecting

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microorganisms.26-29 It also has an accompanying ozone-based in situ compatible lysis technique.30, 31 More

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importantly, we have recently demonstrated that the NanoGene assay is as a viable method for detection of

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Microcystis aeruginosa (M. aeruginosa).32

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Using M. aeruginosa as the target cyanobacterium, we evaluated the performance of the

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electrocoagulator with both laboratory and environmental samples (i.e., river water with a range of

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eutrophication). Cell densities ranged from 1.7 × 105 to 4.1 × 106 and 6.5 × 103 to 6.6 × 107 cells·mL-1,

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respectively. M. aeruginosa was chosen because the genus Microcystis is abundant and widespread in freshwater

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bodies.33,

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measurement were employed to evaluate the pre-concentration efficiency for both varying cell densities and

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operation duration. The NanoGene assay was used to quantify the microcystin synthetase D, or mcyD gene, in

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both laboratory and environmental samples that were pre-concentrated by the electrocoagulator. The results were

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compared to those pre-concentrated by a centrifuge. Zeta potential measurements were also performed to further

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elucidate the degree of pre-concentration in the environmental samples. In this way, we could establish the

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suitability of the electrocoagulator as an in situ compatible alternative to the centrifuge for cyanobacterial

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detection.

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The absorbance ratio (before and after pre-concentration), as well as the dry cell weight

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EXPERIMENTAL SECTION

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Design and operation of the electrocoagulator. The electrocoagulator consisted of a pair of electrolysis

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electrodes, a modified vial, trapping gauze and aspiration outlet (Figure 1a and 1b). The electrodes were 5



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commercial off-the-shelf aluminum tubing (KS Aluminum Tubing #8100, Chicago, Illinois, USA) cut into

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lengths of 60 mm. They were suspended (3.5 mm apart with a silicone spacer) inside a 4-mL modified vial with

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an aspiration outlet at its base. The overall size of the electrocoagulator is ~2 × 3 × 10 cm with a weight of ~16 g

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(without battery), and the materials cost less than 3 US dollars.

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Prior to operation, the trapping gauze (a loosely woven cloth with average pore size > 0.2 mm as shown

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in the insert of Figure 1b) was secured over the aspiration outlet. During operation, 4.5 VDC (~70 mA, 300 mW)

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was applied between the electrolysis electrodes to enable electrocoagulation of cyanobacteria in the sample.

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Note that the electrical power may be supplied by three AA size batteries. Gases generated from the resulting

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electrolysis were allowed to vent freely. After operation, a syringe was used to manually aspirate the culture

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broth from the electrocoagulator via the aspiration outlet. In this way, the electrocoagulated cyanobacteria were

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collected on the trapping gauze and this completed the pre-concentration of the samples. The pre-concentrated

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cyanobacteria from the gauze were subjected to genomic DNA (gDNA) extraction for the NanoGene assay. The

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pre-concentrated cyanobacteria from the gauze were subjected to genomic DNA (gDNA) extraction for the

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NanoGene assay. The captured cyanobacteria were first extracted from the gauze by a tweezer. The remaining

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cyanobacteria on the gauze was further extracted by soaking it in the lysis buffer (DNA extraction kit). It is

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important to note that unlike conventional filtration using a fine membrane, a pump with high operating head is

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not required. In this case, manual aspiration of the culture broth via a syringe sufficed and this was only possible

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due to the large pore size of the trapping gauze.

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Laboratory sample preparation. The target cyanobacterium M. aeruginosa strain UTEX 2388 was obtained

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from the Culture Collection of Algae at the University of Texas, Austin, USA. It was inoculated into a 250-mL

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Erlenmeyer flask containing 100 mL modified Bold 3N medium in which soil water was eliminated. The flask

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was subsequently incubated at ambient temperature with shaking at 100 rpm to facilitate aeration. Continuous 6


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illumination at 20,000 lux (30 W, SL230D, City E.L.G., Incheon, Korea) was also employed. M. aeruginosa

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culture broth in early exponential phase was used for the laboratory samples in subsequent experiments.

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The calibration curve (Figure S1) of laboratory sample cell density (cells·mL-1) versus absorbance (as

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optical density or OD) at 680 nm was obtained via a counting chamber under a light microscope and

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spectrofluorometer (SpectraMax M2, Molecular Devices, Sunnyvale, USA), respectively.

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Environmental sample sites and sample preparation. Environmental samples were obtained from 2 negative

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control sites (NC1 and NC2) and 4 study sites (S1, S2, S3 and S4) from the Han River, Korea, in August 2016

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(Figure 2). Han River is a major river of South Korea that has a river basin of 26,018 km2 of basin and 514.8 km

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of stems, respectively. Together they supply water to more than 10 million people in the Seoul metropolitan area.

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The area is also prone to recurring harmful algal blooms.35, 36

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As shown in Figure 2b and c, the waters in the NC1 and NC2 negative control sites were clear and

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sedimentary bottom was visible. On the other hand as shown in Figure 2d, e, f and g, the waters of the S1, S2,

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S3, and S4 study sites were increasingly turbid and a harmful algal bloom was visible in both study sites S3 and

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S4.

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Sampling was performed during hot weather with an average water temperature of ~29ºC. No rainfalls

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was reported during the sampling period. At each sampling site, 2 L of water was drawn from a depth of 30 cm.

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The water quality data for the environmental negative control and test samples are given in the Supplementary

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Information (Table S1). The dominant algal species in the samples were identified via microscopic examination

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(Table 1). The genus Microcystis was predominant in samples S3 and S4.

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Environmental positive control samples (PC1 and PC2) were prepared by spiking M. aeruginosa culture

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broth into the NC1 and NC2 samples to achieve final cell densities of 7.4 × 105 cells·mL-1 (equivalent to OD680

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nm =

0.41) and 6.8 × 105 cells·mL-1 (equivalent to OD680 nm = 0.39), respectively. 7



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Cell densities of environmental samples were determined by counting cells with a counting chamber

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under a light microscope (Figure S2a). The corresponding chlorophyll-a concentration for environmental

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samples is shown in Figure S2b.

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Comparison of electrocoagulator and centrifuge. The samples (laboratory and environmental) in aliquots of 3

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mL were first transferred to the modified 4-mL vial of the electrocoagulator. After electrocoagulation and

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aspiration of the culture broth, the trapping gauze was removed from the electrocoagulator. Subsequently, the

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electrocoagulated sample on the trapping gauze was dried at 80ºC overnight for dry cell weight measurement. In

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order to visualize the samples before and after electrocoagulation, additional aliquots of 3 mL were also

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electrocoagulated in cuvettes separately.

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Similarly, 3-mL test samples were transferred to the 15-mL centrifuge tubes. The centrifuge (Model

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1248, LaboGene, Seoul, Korea) was operated at 2,500 rpm (1,224 relative centrifugal force or RCF) for a given

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duration. The supernatant was then discarded, and the centrifuged pellet in the tubes was also dried at 80ºC

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overnight.

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Afterwards, the gram-dry cell weight per liter (g-DCW·L-1) of the sample was determined by weighing

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the pellet. The pre-concentration efficiency was estimated via the culture broth’s absorbance at 680 nm before

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(ODt0) and after (ODt1) pre-concentration as follows:

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Pre-concentration Efficiency (%) =

× 100

The background absorbance (culture broth only) was subtracted from both ODt0 and ODt1.

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The experiments were performed at 4.5 V for varying operation durations from 0 to 300 s at a cell

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density of 4.1 × 105 cells·mL-1, where OD680 nm = ~0.27. They were also performed for varying cell densities

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from 1.7 × 105 to 4.1 × 106 cells·mL-1, where OD680 nm ranged from 0.08 to 1.1, with an operation duration of

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180 s. All experiments were performed in biological triplicates unless otherwise stated. Note that uncoagulated 8


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laboratory samples were also subjected to culture broth aspiration in the electrocoagulator. This was to establish

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the inability of the trapping gauze to capture cyanobacteria without electrocoagulation.

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Analysis of pre-concentrated samples via NanoGene assay. Briefly, the NanoGene assay consisted of

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magnetic beads (MB) and dual quantum dot nanoparticles (QD).26-28, 32 The aminated MB (2 × 107 beads·mL-1,

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Dynabead M270, Invitrogen, Carlsbad, USA) were coupled with carboxyl QD565 (2 µmole·L-1, Invitrogen,

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Carlsbad, USA) through an amide bond formation to form the MB-QD565 conjugate. It was further conjugated

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with the probe DNA (100 µmole·L-1, Bioneer, Daejeon, Korea) to form the MB-QD565-probe DNA complex. The

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signaling DNA (100 µmole·L-1, Bioneer) was separately immobilized on the surface of QD655 (2 µmole·L-1,

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Invitrogen, Carlsbad, USA) nanoparticles to form the signaling DNA-QD655 complex. During DNA

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hybridization, the MB-QD565 complex was tethered to the signaling DNA-QD655 complex via the target mcyD

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gene through complementary base pairing. After rinsing to remove untethered complexes, the target mcyD gene

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could be quantified by the normalized fluorescence of QD655 with respect to that of QD565.

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The pre-concentrated samples (laboratory and environmental) from both the electrocoagulator and

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centrifuge were first subjected to gDNA extraction. This was performed in duplicate using the NucleoSpin Plant

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II DNA extraction kit (Macherey-Nagel, Düren, Germany) in accordance with the manufacturer’s instructions.

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The extracted gDNA was eluted in 100 µL of elution buffer, and duplicate gDNA samples were pooled together.

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The concentration of the pooled gDNA extracts was spectrophotometrically measured at OD260

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NanoDropTM 2000 (Thermo Fisher, Wilmington, USA). The gDNA extracts were then denatured at 95ºC for 20

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min prior to DNA hybridization.

nm

using a

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DNA hybridization was initiated by adding 5 µL of denatured gDNA extracts to 300 µL of DIG Easy

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Hybridization buffer (Roche, Basel, Switzerland). The mixture also contained MB-QD565-probe DNA and

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QD655-signaling DNA complexes. This was followed by the incubation at 37ºC for 15 h with a gentle tilt rotation. 9



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After incubation, the sample was rinsed 3 times with 0.1 mole·L-1 phosphate buffer (pH 7.4). The fluorescence

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intensities of QD565 and QD655 were measured on black 96-microplates (Thermo Fisher Scientific) using a

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SpectraMax M2 spectrofluorometer with emission wavelengths at 570 and 660 nm, respectively, and an

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excitation wavelength of 360 nm. Normalized fluorescence was calculated as the ratio of measured fluorescence

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QD655/QD565.

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The negative control for the NanoGene assay was in the form of ultrapure deionized water

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(DNase/RNase/Protease free, Intron Biotechnology, Gyeonggi, Korea) added to the DNA hybridization reaction

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instead of denatured gDNA.

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A standard curve for the NanoGene assay (Figure S3) was established using the amplified mcyD gene

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fragment (297 bp) of M. aeruginosa (mcyD gene copy numbers ranging from 6.5 × 100 to 6.5 × 1010). It was

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prepared by PCR using the gDNA of M. aeruginosa and the primer set of mcyDF2 and mcyDR2.37 The PCR

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mixture and thermocycling conditions were described previously in detail.31 The mcyD gene copy numbers were

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calculated using Avogadro’s number (i.e., 6.022 × 1023 molecules·mole-1) and the DNA weight in Daltons, with

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the assumption that the average weight of a base pair (bp) is 650 Daltons. The limit of quantification for the

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NanoGene assay was 7 mcyD gene copy number·mL-1 (Figure S3).

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Zeta potential measurement. Zeta potential measurement was performed by laser Doppler velocimetry using

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the zeta potential and particle size analyzer ELSZ-2000 (Otsuka Electronics, Osaka, Japan). The zeta potential (ζ)

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of the environmental samples was measured before and after pre-concentration (by electrocoagulator and

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centrifuge). Three milliliters of the environmental sample was transferred to the zeta flow cells (Otsuka

218

Electronics) and measured. Electrophoretic analysis was carried out in technical triplicates.

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RESULTS AND DISCUSSION 10


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Pre-concentration and NanoGene assay analysis of laboratory samples. Using the electrocoagulator in the

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absence of applied voltage at the electrolysis electrodes, there was no electrocoagulation of cyanobacteria. As

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expected, no cyanobacteria were observed on the trapping gauze after aspiration (Figure 3a). This means that in

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the absence of electrocoagulation, the cyanobacteria in the laboratory samples were sufficiently small to simply

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pass through the trapping gauze with near zero percent pre-concentration efficiencies for cell densities ranging

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from 1.4 × 105 to 4.1 × 106 cells·mL-1 (Figure 3b). This is because the size of individual M. aeruginosa cells was

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too small (~10 to 40 µm) to be captured by the trapping gauze (average pore size > 0.2 mm). However, with

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operation durations of 60 s and higher, initial visual observations suggested that it was possible to pre-

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concentrate the laboratory samples for the given range of cell densities (Figure 3c and 3d). A visible increase in

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the amount of cyanobacterial cells captured by the trapping gauze could be seen with an increase in operation

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duration (60 – 300 s).

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With varying operation durations (Figure 4a and 4b), both the pre-concentration efficiency and dry cell

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weight resulting from the electrocoagulator were slightly lower if not comparable to that of the centrifuge (cell

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density at 4.1 × 105 cells·mL-1). At an operation duration of 180 s, the electrocoagulator and centrifuge achieved

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similar pre-concentration efficiencies of 59.1 ± 3.7% and 61.9 ± 2.6% (p-value = 0.342, as determined by a t-test,

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Table S2), respectively. Their corresponding dry cell weights were also near identical at 0.14 ± 0.02 and 0.14 ±

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0.05 g-DCW·L-1 (p-value = 1.000, Table S2), respectively. Increasing the operation duration of the

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electrocoagulator from 180 to 300 s of the electrocoagulator resulted in a marginal increase in pre-concentration

239

efficiency to 62.8 ± 9.9% (~3% increase). Therefore, the nominal operation duration of the electrocoagulator for

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subsequent experiments was set at 180 s.

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With varying cell densities (Figure 4c and 4d), both the pre-concentration efficiency and dry cell weight

242

from the electrocoagulator were also slightly lower if not comparable to that of the centrifuge (nominal

243

operation duration of 180 s). For cell densities from 3.2 × 105 to 4.1 × 106 cells·mL-1, the pre-concentration 11



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efficiencies of electrocoagulator appeared to be stable at ~60% which was consistent with the earlier experiment.

245

They were slightly lower than that of the centrifuge (~70%).

246

At lower cell densities (2.2 × 105 and 1.7 × 105 cells·mL-1), the pre-concentration efficiencies decreased

247

to 40.4 ± 5.4 and 33.9 ± 0.2%, respectively. This trend was also observed with the centrifuged samples where the

248

pre-concentration efficiencies also decreased to 53.1 ± 0.9 and 37.5 ± 1.8% for these cell densities, respectively.

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As shown in Figure 4e and 4f, the pre-concentrated samples from the electrocoagulator were analyzed

250

successfully with the NanoGene assay. As expected, the normalized fluorescence and mcyD gene copy number

251

increased with cell density for both the electrocoagulator and centrifuge. At 2.5 × 106 cells·mL-1, pre-

252

concentration via electrocoagulator and centrifuge yielded comparable normalized fluorescence of 4.49 ± 0.46

253

and 5.04 ± 0.51, respectively. The mcyD gene copy numbers from the electrocoagulator and centrifuge were not

254

significantly different (p-value > 0.05, as determined by a t-test, Figure 4f and Table S2).

255 256

Pre-concentration of environmental samples. As expected, using the electrocoagulator and centrifuge (Figure

257

5a and 5b) both yielded pre-concentrated cyanobacteria in PC1 and PC2 (positive control) but not in NC1 and

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NC2 (negative control). There were also no visible pre-concentrated cyanobacteria obtained from S1 and S2.

259

However, the electrocoagulator appeared to be able to pre-concentrate S3 and S4 as well as, if not more

260

effectively, than the centrifuge. As shown in the dotted boxes in Figure 5a and 5b, S3 formed a well-defined

261

layer after electrocoagulation while remained dispersed after centrifugation. Similarly, the electrocoagulator was

262

able to form a well-defined layer of cyanobacteria in S4. More specifically, the electrocoagulator was able to

263

pre-concentrate S3 (at 57.8 ± 5.1%) with corresponding dry cell weight of 0.33 ± 0.06 g-DCW·L-1 while the

264

centrifuge was unable to do so (Figure 6, p-value ± 30 mV indicate well dispersed particles with no

285

aggregation.42 After pre-concentration by the electrocoagulator and centrifuge, their zeta potentials were reduced

286

as expected. Similarly, the zeta potential for S4 was reduced after pre-concentration, but by a lesser amount than

287

expected. This could be due to the 10-fold dilution that was necessary for zeta potential measurement. With the

288

electrocoagulator, the zeta potential of S3 was reduced significantly from -16.51 ± 1.02 to -5.05 ± 1.05 mV (p13



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value = 0.008, Table S4). However, with the centrifuge, it was marginally reduced to -14.32 ± 0.86 mV (p-value

290

= 0.333, Table S4). Therefore the electrocoagulator was able to pre-concentrate S3 more effectively than the

291

centrifuge.

292 293

NanoGene assay analysis of pre-concentrated environmental samples. With reference to Figure 7, M.

294

aeruginosa in the environmental samples pre-concentrated by the electrocoagulator were successfully quantified

295

via the NanoGene assay. The normalized fluorescence for S3, S4, PC1 and PC2 was 3.0 ± 0.7, 4.8 ± 0.6, 4.2 ±

296

0.3 and 4.3 ± 0.3 (Figure 7a), and these corresponded to 3.5 × 103, 5.6 × 106, 1.2 × 105, and 2.0 × 105 mcyD gene

297

copy number (Figure S3 and 7c). More importantly, it demonstrates that the electrocoagulator is also compatible

298

with the NanoGene assay for environmental samples, as it was for the laboratory samples shown earlier.

299

However, with the centrifuge, the normalized fluorescence for S4, PC1 and PC2 was 4.4 ± 0.3, 4.8 ± 0.2 and 4.3

300

± 0.6 (Figure 7b) and these corresponded to 2.6 × 105, 1.2 × 106, and 5.1 × 105 mcyD gene copy number (Figure

301

7d).

302

The viability of the electrocoagulator as an alternative to the centrifuge could be further highlighted by

303

comparing its NanoGene analysis results to the environmental samples’ cell density (Figure S2a) and

304

chlorophyll a concentrations (Figure S2b). The environmental samples’ cell density for S1, S2, S3 and S4 were

305

8.6 × 103, 6.5 × 103, 2.7 × 105 and 6.6 × 107 cells·mL-1. Both the electrocoagulator and centrifuge were not able

306

to pre-concentrate S1 and S2 (cell density less than 104 cells·mL-1). However the electrocoagulator was able to

307

pre-concentrate both S3 and S4. On the other hand, the centrifuge was only able to pre-concentrate S4.

308

The successful NanoGene assay analysis of the environmental samples pre-concentrated by the

309

electrocoagulator verified the compatibility of these technologies. The electrocoagulator has been shown to be

310

an in situ compatible alternative to the centrifuge. More importantly, the electrocoagulator is also small, light

311

weight and low cost. In accordance with the World Health Organization 2003 guidelines for safe recreational 14


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water environments, the operational cell density limit of the electrocoagulator at ~105 cells·mL-1 also

313

corresponded to the moderate probability of adverse health effects.43 This means the electrocoagulator can be

314

used on environmentally relevant concentration of cyanobacteria. In other words, the electrocoagulator has

315

advanced the possibility of in situ cyanobacterial detection in eutrophic waters by reducing the size, weight and

316

cost associated with pre-concentration of cyanobacteria.

317 318

ASSOCIATED CONTENT

319

Supporting Information

320

The supporting information is available free of charge on the ACS Publication website. This includes cell

321

density of laboratory cultures (Microcystis aeruginosa), chlorophyll-a concentrations of environmental samples,

322

the calibration curve of the NanoGene assay, and water quality data of environmental test samples.

323 324

ACKNOWLEDGEMENTS

325

This work was supported by the National Research Foundation of Korea (NRF-2017R1A2B4005133 and NRF-

326

2015R1D1A1A01060317).

327

15



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REFERENCES

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(41) Komárek, J. Coccoid and colonial cyanobacteria A2 - WEHR, JOHN D. In Freshwater Algae of North

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FIGURE CAPTIONS

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Figure 1. (a) Schematic of the portable electrocoagulator for in situ cyanobacterial detection with the NanoGene

430

assay. (b) Photo of the electrocoagulator with insert showing the trapping gauze.

431

Figure 2. (a) Sampling site map. Negative control sites (b) NC1 (c) NC2. Study sites (d) S1 (e) S2 (f) S3 (g) S4.

432

Figure 3. (a) Photo of the trapping gauze when used with the uncoagulated laboratory samples. (b)

433

Experimental plot of the pre-concentration efficiency (%) versus cell density (cells·mL-1) for uncoagulated

434

laboratory samples for varying cell densities (1.4 × 105 to 4.0 × 106 cells·mL-1). (c) Laboratory samples (4.1 ×

435

105 cells·mL-1) pre-concentrated via the electrocoagulator with varying duration 0 – 300 s at 4.5 VDC. (d)

436

Laboratory samples of varying cell densities (1.7 × 105 to 4.1 × 106 cells·mL-1) pre-concentrated via the

437

electrocoagulator at 4.5 VDC for duration of 180 s.

438

Figure 4. Pre-concentration and dry cell weight of laboratory samples via electrocoagulator and centrifuge for

439

(a)(b) varying operation durations at a cell density of 4.1 × 105 cells·mL-1 and (c)(d) varying cell densities with

440

an operation duration of 180 s. (e)(f) NanoGene assay analysis of laboratory samples pre-concentrated using

441

electrocoagulator and centrifuge.

442

Figure 5. Photos of environmental samples before and after pre-concentration using (a) electrocoagulator and (b)

443

centrifuge. Pre-concentration of S3 and S4 (eutrophic water samples) was more effective using the

444

electrocoagulator as compared to the centrifuge (dotted box).

445

Figure 6. (a)(b) Pre-concentration efficiency and (c)(d) dry cell weight of environmental samples via

446

electrocoagulator and centrifuge. ND stands for non-detectable.

447

Figure 7. NanoGene assay analysis of pre-concentrated environmental samples via electrocoagulator and

448

centrifuge. (a)(b) Normalized fluorescence (QD655/QD565) and (c)(d) mcyD gene copy numbers. ND stands for

449

non-detectable.

450 21



(a)

(b)

Figure 1

22


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(a) Sampling Site Map

(b) NC1

(c) NC2

(d) S1

(e) S2

(f) S3

(g) S4

Figure 2 23



(a)

(b)

(d)

(c)

Figure 3

24


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(a)

(b)

(c)

(d)

(e)

(f)

Figure 4

25



(a)

(b)

Figure 5

26


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(a)

(b)

(c)

(d)

Figure 6

27



(a)

(b)

(c)

(d)

Figure 7

28


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Table 1. Dominant species of cyanobacteria in environmental test samples.

Environmental samples

Dominant algal species

NC1

ND

NC2

ND

S1

Euglena

S2

Scenedesmus, Euglena

S3

Microcystis, Chlamydomonas, Eudonina, Euglena, Spirulina, Anabaena, Closterium, Cosmarium

S4

Microcystis, Chlamydomonas, Euglena, Cosmarium

ND denotes non-detectable

29



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Table 2. Zeta potential measurements of environmental samples before and after pre-concentration (via electrocoagulator and centrifuge).

Environmental samples

Raw water sample (mV)

Electrocoagulator (mV)

Centrifuge (mV)

NC1

6.63 ± 1.45

NA

NA

NC2

6.58 ± 0.21

NA

NA

S1

6.46 ± 0.73

NA

NA

S2

4.94 ± 1.31

NA

NA

S3

-16.51 ± 1.02

-5.05 ± 1.05

-14.32 ± 0.86

S4a

-21.48 ± 1.20

-16.97 ± 2.47

-17.65 ± 1.64

PC1

-32.18 ± 0.03

-19.99 ± 0.29

-20.55 ± 1.38

PC2

-35.09 ± 0.54

-27.83 ± 0.42

-23.54 ± 0.31

Values represent the mean ± SD (n = 3). NA denotes not applicable. S4a sample was ten-fold diluted with distilled water.

30


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Electrocoagulation as pre-concentration technique for cyanobacterial detection by NanoGene assay.


Detection of cyanobacteria in eutrophic water using a portable

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