Polyphasic toxicological screening of Cylindrospermopsis raciborskii and Aphanizomenon gracile isolated in Poland

Algal Research 24 (2017) 72–80

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Polyphasic toxicological screening of Cylindrospermopsis raciborskii and Aphanizomenon gracile isolated in Poland Piotr Rzymski a,⁎, Barbara Poniedziałek a, Joanna Mankiewicz-Boczek b,d, Elisabeth J. Faassen c, Tomasz Jurczak d, Ilona Gągała-Borowska b, Andreas Ballot e, Miquel Lürling c, Mikołaj Kokociński f a

Department of Environmental Medicine, Poznan University of Medical Sciences, Poznań, Poland European Regional Centre for Ecohydrology of the Polish Academy of Sciences, Łódź, Poland Department of Environmental Sciences, Wageningen University, Wageningen, The Netherlands d Department of Applied Ecology, Faculty of Biology and Environmental Protection, University of Łódź, Łódź, Poland e Norwegian Institute for Water Research, Oslo, Norway f Department of Hydrobiology, Adam Mickiewicz University, Poznań, Poland b c

a r t i c l e

i n f o

Article history: Received 5 October 2016 Received in revised form 17 February 2017 Accepted 21 February 2017 Available online 22 March 2017 Keywords: Cylindrospermopsis raciborskii Aphanizomenon gracile Cyanotoxins α-γ,-Diaminobutyric acid In vitro toxicity

a b s t r a c t Aphanizomenon gracile and Cylindrospermopsis raciborskii are extensively studied Nostocales of wide geographical distribution and have potential to produce toxins. However, a number of knowledge gaps regarding their toxicity and related health risks in certain locations, including Europe, exists. The present study applied a polyphasic approach to screen the toxicity of different strains of C. raciborskii (LBY-Cr, LBO-Cr and LKM-Cr) and A. gracile (LBY-Ag, LBN-Ag and LWI-Ag) isolated from five freshwater lakes of Western Poland. The following investigations were carried out: (i) in vitro toxicological studies employing human cells isolated from healthy donors; (ii) analytical screening for the presence of cylindrospermopsin (CYN), guanidinoacetate (GAA; initial CYN precursor and postulated general cyanobacterial metabolite), three microcystin (MC) analogues, β-N-methylaminoL-alanine (BMAA) and its isomer α-γ,-diaminobutyric acid (DAB), anatoxin-a (ATX) and ten saxitoxin (STX) analogues; and (iii) molecular studies of genes involved in CYN, GAA, MCs and ATX biosynthesis. Extracts of C. raciborskii LBY-Cr and A. gracile LBN-Ag caused a significant increase in the intracellular reactive oxygen content in human neutrophils during short-term (1 h) exposure and also led to lipid peroxidation and cell death. No cytotoxic effects were noted for the other tested strains. None of the toxin genes (cyrA, cyrJ, anaF and mcyE) and toxins (CYN, GAA, MCs, BMAA, ATX and STX) were detected. The only exception was DAB found at a concentration below 1.0 μg g−1 dw in A. gracile LWI-Ag. It is the first time that cyanobacterial DAB producer has been identified in the Central European region. The study points to the production of as yet unknown metabolite(s) that may pose a relevant threat to human health through strains of C. raciborskii and A. gracile isolated from two Polish lakes, and adds to the general understanding of the toxicity of European strains of both species. © 2017 Elsevier B.V. All rights reserved.

1. Introduction Cylindrospermopsis raciborskii and members of the traditional “Aphanizomenon” genus are the most studied Nostocales owing to their nearly global distribution that encompasses Africa, Australia, Asia, Europe, North and South America, and to their potential to biosynthesize a wide range of toxic compounds which vary in mechanisms of action [14,61]. Some C. raciborskii strains can produce teratogenic polymethoxy-1-alkenes [33], cytotoxic cylindrospermopsin (CYN) [52], and neurotoxic saxitoxins (STXs) [30,31]. The biosynthesis

⁎ Corresponding author. E-mail address: [email protected] (P. Rzymski).

http://dx.doi.org/10.1016/j.algal.2017.02.011 2211-9264/© 2017 Elsevier B.V. All rights reserved.

of CYN, STXs as well as anatoxin-a (ATX) was also confirmed for “Aphanizomenon” spp. [6,14]. However, the ability to produce certain compounds is highly species- and strain-specific and geographically diversified [15,61]. Furthermore, particular strains may vary in the level of produced toxin, as recently evidenced for CYN synthesis in Australian strains of C. raciborskii [71]. The complex assessment of risks arising from the occurrence of certain cyanobacteria requires well-designed and multi-approach investigations [32]. To date, knowledge gaps regarding the toxicity of nostocaleans still remain and this also concerns important species inhabiting European freshwaters. For example, strains of C. raciborskii from this region have never been found to produce CYN, ATX or STX [36,61], yet their exudates exhibited toxic action in rodents or in vitro experimental models [2,10,25,54]. Aphanizomenon gracile, in turn, has been recognized as a

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main producer of CYN in Polish freshwater lakes [36], some of its strains isolated from German and Mediterranean countries were confirmed to biosynthesize STXs [7,15,38]. The production of some other important compounds such as neurotoxic amino acid β-methylamino-L-alanine (BMAA) and its isomer 2,4-diaminobutyric acid (DAB) has never been assessed for European isolates of C. raciborskii and A. gracile. The determination of these compounds would be of high interest, particularly if one considers that extracts of C. raciborskii strains isolated from different locations in Hungary evoked neurotoxicity involving inhibition of the acetylcholine responses in Helix pomata [69] - this effect has also been observed in rats chronically exposed to BMAA via intracerebroventricular administration [56]. A previous study has already identified BMAA and DAB in cyanobacterial scums collected in The Netherlands [22] although information concerning their potent producers is still scarce. It has been recently evidenced that freshwater cyanobacteria are capable of producing guanidinoacetate (GAA) [8]. As established by Mihali et al. [48], GAA is a precursor of CYN biosynthesis encoded by cyrA (AoaA homolog). However, its production not only by CYN producers but also by CYN-negative species, including Microcystis aeruginosa, suggested that GAA can be a general cyanobacterial metabolite [8]. This finding is important in view of GAA toxicity implicated, i.e., in neural dysfunction through enhancement of acetylcholinesterase activity, the main enzyme that hydrolyses acetylcholine [72]. Further research is required to elucidate how frequently GAA is present in freshwater cyanobacteria species and strains. Moreover, it was demonstrated that some nonCYN producing strains of C. raciborskii revealed the presence of unknown toxic compound(s) that partially mimic the allelopathic CYN action through up-regulation of alkaline phosphatase in phytoplankton [9, 62,63]. One could therefore hypothesize that the CYN biosynthesis pathway may be initiated in these strains but terminated at some earlier step, and that resulting CYN intermediates are responsible for the reported effect. The analytical screening of GAA, preferably coupled with cyrA studies, would be helpful in establishing whether or not this is the case. As demonstrated, some CYN-negative strains of Chrysposporum (Aphanizomenon) ovalisporum and Cuspidothrix issatschenkoi can contain cyr cluster genes (cyrA, cyrB, cyrC) although the production of encoded products has not been assessed [5,66]. The present research was undertaken to compare the toxicities of C. raciborskii and A. gracile strains isolated from freshwater lakes of Western Poland. A polyphasic approach was implemented and included (i) investigations of the response of human neutrophils isolated from healthy donors to cyanobacterial extracts using three complimentary assays, (ii) analytical determinations of ATX, microcystins (MCs), CYN, GAA, DAB, BMAA and STXs, and (iii) screening for selected genes known to be involved in the production of ATX, CYN, GAA and MC in cyanobacteria. We hypothesized that: (i) exudates of all investigated strains would impact human neutrophils, (ii) all strains contain GAA, and (iii) other cyanotoxins are present depending on corresponding genes presence/absence in the strains studied. 2. Materials and methods 2.1. Cyanobacteria strains, isolation and culture The study evaluated the toxicities of three strains of C. raciborskii and three strains of A. gracile isolated from freshwater lakes located in Western Poland. C. raciborskii strains were isolated from Lake Boczowskie (52°19′10″N, 14°56′47″E), Lake Bytyńskie (52°29′55″N, 16°30′30″E) and Lake Kierskie Małe (52°29′12″N, 16°47′14″E), and named LBO-Cr, LBY-Cr and LKM-Cr, respectively. A. gracile strains were isolated from Lake Bytyńskie and Lake Bnińskie (52°12′02″N, 17°06′59″E) and Lake Witobelskie (52°15′55″N, 16°43′32″E) and named LBY-Ag, LBN-Ag and LWI-Ag, respectively. The morphometric parameters and detailed physico-chemical conditions of the lakes can be found in [36]. All strains were isolated under a Carl Zeiss light and inverted microscopes (model

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Axioplan and Primo Vert, respectively) (magnification 400×) from collected water samples. Single filaments were isolated using elongated glass Pasteur pipette and transferred to culture flasks containing sterile BG-11 media (Sigma–Aldrich, USA). This procedure was repeated until monocultures of the cyanobacteria were obtained. The isolates were incubated in 250 mL Erlenmeyer flasks containing 150 mL of sterile BG-11 medium in an incubation chamber (POL-EKO, Poland) at 21 °C under 80 μmol m−2 s−2 irradiance using cool white fluorescent light with a photoperiod regime of 12 h dark and 12 h light. The strains were maintained in the culture collection of the Department of Hydrobiology at Adam Mickiewicz University. 2.2. Taxonomic determination The molecular analysis was performed using cyanobacterial gene fragment of 16S rRNA, with the primer pair Cyano 16S F (forward): CGGACGGGTGAGTAACGCGTG, and Cyano 16S R (reverse): CCCATTGCGGAAAATTCCCC. The primer set amplifies a PCR product of 258 bp [39]. The amplification reaction was carried out for the each strain in a volume of 30 μL mix containing 50–200 ng of DNA template, 1 × PCR buffer, 3 mM MgSO 4 , 0.2 mM dNTPs, 0.5 μM of each primer, 0.1 mg mL − 1 BSA and 1.25 U of Pfu Taq polymerase (Thermo Scientific, USA). The PCR program, performed in an Eppendorf MasterCycler Gradient thermocycler, started with an initial denaturation at 95 °C for 10 min. Subsequently, 26 cycles of denaturation at 94 °C for 10 s, primer annealing at 58 °C for 30 s, and an extension at 70 °C for 1 min. Prior to sequence analysis, the PCR products were separated and excised from a 1.0% agarose gel bathed in ethidium bromide and purified with the QIAEX II® Gel Extraction Kit according to the manufacturer's instructions. Purified products were sequenced by Genomed® laboratory in Warsaw, Poland (http://www.genomed.pl/). Assembled DNA sequences were edited and aligned using the Bioedit Sequence Alignment Editor (Ibis Biosciences Carlsbad, USA). A Basic Local Alignment Search Tool (BLAST) was used to verify gene homology in the GenBank database (National Institute of health, USA). 2.3. Extract preparation Cell-free extracts of A. gracile and C. raciborskii strains were prepared by subjecting cell suspensions containing 55,000 individuals per mL to ultrasonic treatment on ice (2 min in 2 cycles with 60 s break). Complete cell lysis was confirmed by microscopic examination. The broken cell suspensions were centrifuged (12,000g, 10 min). The supernatants were sterilized by filtration on 0.22 μm syringeless filter devices (Carl Roth, Germany) and stored at −20 °C until use. 2.4. In vitro toxicity studies The present study evaluated the toxic effects of C. raciborskii and A. gracile extracts in human-derived neutrophils, the most abundant and chemo-sensitive leukocytes with a circulating half-life of only 6–8 h [51], previously shown as a convenient in vitro model to evaluate the toxicity of cyanobacterial metabolites [54,64]. The employed model is rapid in contrast to those employing cell lines that usually require culturing and a longer assay time to allow for cell responses. Heparinized blood samples (6.0 mL), collected in lithium heparin tubes from 5 healthy (screened by physical examination, medical history and initial blood tests), non-smoking and normal weighted (BMI 18.5–24.9) donors (aged 21–25 years old; 2 female, 3 male) was purchased from the Regional Centre of Blood and Blood Treatment in Poznan, Poland, according to accepted safeguard standards and legal requirements in Poland. Human neutrophils were isolated using a one-step density-gradient centrifugation on Gradisol G of a specific gravity of 1.115 g mL−1 (Polfa, Poland) at 400 g at room temperature for 30 min. The residual erythrocytes were removed from the cell

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population by hypotonic lysis. The purity of the neutrophils (N90%) was verified by counting under a light microscope after May-GrunwaldGiemsa staining. Using three different assays, the viability of cells, intracellular generation of reactive oxygen species (ROS) and lipid peroxidation were assessed in neutrophils exposed to cyanobacterial extracts for 1 h. Each experimental assay was performed on five independent neutrophil isolates obtained from the blood of five different healthy donors. 2.4.1. Intracellular reactive oxygen species assay Human neutrophils were loaded for 30 min at 37 °C in darkness with 20 μM of 2′,7′–dichlorofluorescin diacetate (DCFDA; Abcam, UK), a fluorogenic dye that measures hydroxyl, peroxyl and other reactive oxygen species (ROS) activity within the cell. Cells were then washed, dissolved in phosphate-buffered saline (PBS) and seeded in a black clear bottom 96-well plate at a density of 20 × 104 neutrophils per well (90 μL aliquots/well). Cyanobacterial extracts were added to the amount of 10 μL each, and the plate was incubated at 37 °C for 1 h. Fluorescence of DCFDA was measured kinetically after 15, 30 and 60 min of incubation using a Synergy HTX multi-mode plate reader (BioTek, USA) at an excitation of 495 nm and emission of 528 nm. The background signal, measured in exposed neutrophils not loaded with DCFDA, was substrated from the corresponding samples. Three technical replicates were conducted for each donor. The final results were presented as a percentage of the parallel control constituted of neutrophils incubated with 10 μL of PBS. 2.4.2. Lipid peroxidation assay Lipid peroxidation was analyzed using a Lipid Peroxidation Colorimetric/Fluorometric Assay Kit (BioVision, UK) by means of malondialdehyde (MDA) content. Human neutrophils were seeded in a 96-well plate at a density of 20 × 104 neutrophils per well (90 μL aliquots/well) and exposed to 10 μL of each cyanobacterial extract for 1 h at 37 °C. The control was constituted of cells incubated with 10 μL of PBS. After the experiments, cells were harvested from each well and homogenized on ice in 300 μL of provided lysis buffer (with addition of butylated hydroxytoluene to prevent artificial lipid peroxidation) and centrifuged to remove insoluble material. The resulting 200 μL of supernatants were transferred to a microcentrifuge tube and supplemented with 600 μL of thiobarbituric acid (TBA) to generate an MDA–TBA adduct. To accelerate the reaction, samples were incubated at 95 °C for 60 min and the final product was measured colorimetrically at 532 nm. Three technical replicates were conducted for each donor. The calculated values were compared to a calibration curve prepared using MDA standard (BioVision, UK). The coefficient of variation (r2) for the calibration curve was 0.99. The final results were presented as a percentage of parallel control constituted of neutrophils incubated with 10 μL of PBS. 2.4.3. Cell viability assay Cell viability was determined using the colorimetric 3-(4,5-dimethylthiazol-2-yl)-2,5 diphenyltetrazolium bromide (MTT) metabolic activity assay (BioVision, UK) according to the manufacturer's instructions. Human neutrophils were seeded in a 96-well plate at a density of 20 × 104 neutrophils per well (90 μL aliquots/well) and exposed to 10 μL of each cyanobacterial extract for 1 h at 37 °C. Afterwards, cells were washed, seeded again, and 10 μL of MTT was added to each well for 2 h. Neutrophils were then treated with 10% sodium dodecyl sulfate in 0.01 M HCl and incubated for another 2 h in the darkness. The optical density (OD) of the final product (the formazan crystals) was measured at 570 nm using a Synergy HTX microplate reader (BioTek, USA). Three technical replicates were conducted for each donor. The final results were presented as a percentage of parallel control constituted of neutrophils incubated with 10 μL of PBS.

2.5. Cyanotoxin analyses 2.5.1. Determination of guanidinoacetate Analyses of GAA in cyanobacterial extracts were performed using high-performance liquid chromatography (HPLC; Express LC-100, Eksigent, United States) coupled with a 3200 QTRAP triple-quadrupole linear ion trap mass spectrometer fitted with a TurboIonSpray interface (Applied Biosystems/MDS Sciex, Germany). The chromatographic method was carried out on a Velocity C18-2, 50 × 21 m column (Bionacom, UK), applying a mobile-phase gradient. The conditions of determination followed those previously applied by Barón-Sola et al. [8]. A calibration curve was obtained using a commercial GAA standard (Sigma Aldrich, Germany) dissolved in 10 mM of ammonium acecate. The limit of GAA detection was 0.01 μg mL−1. 2.5.2. Determination of cylindrospermopsin and anatoxin-a To determine CYN and ATX, cyanobacterial extracts were concentrated to dryness by an evaporation process in an SC110A Speedvac® Plus, ThermoSavant (Holbrook, NY, USA). Samples were redissolved in 500 μL of distilled water and filtered through a Gelman GHP Acrodisc 13 mm syringe filter with a 0.45 μm GHP membrane and minispike outlet (East Hills, NY, USA). Chromatographic separation was performed using an Agilent (Waldbronn, Germany) 1100 series HPLC system consisting of a degasser, a quaternary pump, a column compartment thermostat set at 40 °C, and a diode array detector operated at 200– 300 nm on a Merck (Darmstadt, Germany) Purospher STAR RP-18e column (55 mm × 4 mm I.D. with 3 μm particles) protected by a 4 mm × 4 mm guard column. The mobile phase consisted of water (solvent A) and acetonitrile (solvent B), both containing 0.05% trifluoroacetic acid. The flow rate was 1.0 mL min−1 with the following linear gradient programme: 0 min, 1% B; 5 min, 7% B; 5.1 min, 70% B; 7 min, 70% B; 7.1 min, 1% B; stop time, 12 min. The injection volume was 20 μL. Cyanotoxins in the samples were identified by comparing the retention time and UV spectrum (200–300 nm) with an absorption maximum at 262 nm for CYN and 227 nm for ATX. 2.5.3. Determination of microcystins To analyze MC content, cyanobacterial extracts were concentrated as described for CYN and ATX. Prior to HPLC analyses, samples were redissolved in 500 μL of 75% aqueous methanol and filtered in the same manner as for CYN. Chromatographic separation was performed using an Agilent (Waldbronn, Germany) 1100 series HPLC system consisting of a degasser, a quaternary pump, a column compartment thermostat set at 40 °C, and a diode array detector operated at 200– 300 nm on a Merck (Darmstadt, Germany) Purospher STAR RP-18e column (250 mm × 4 mm I.D. with 5 μm particles) protected by a 4 mm × 4 mm guard column. The mobile phase consisted of water (solvent A) and acetonitrile (solvent B), both containing 0.05% trifluoroacetic acid. The flow rate was 0.75 mL min−1 with the following linear gradient programme: 0 min, 30% B; 10 min, 35% B; 40 min, 70% B; 42 min, 100% B; 44 min, 100% B; 46 min, 30% B; stop time, 60 min The injection volume was 20 μL. The content of microcystin-LR (MC-LR), microcystinYR (MC-YR) and microcystin-RR (MC-RR) in the samples was analyzed by comparing the retention time and UV spectrum (200–300 nm with an absorption maximum at 238 nm). 2.5.4. Determination of saxitoxins STX determination was performed by liquid chromatography-tandem mass spectrometry (LC-MS/MS), based on [57], but without a solid-phase extraction step prior to analysis. Cyanobacterial culture (85–100 mL) was collected on a GF/C filter and lyophilized. Filters were thrice extracted for 10 min in 0.1 M HCl at 95 °C. After drying the extract down in a speedvac (Thermo Scientific Savant SPD121P, USA), samples were reconstituted in 800 μL of 20 mM HCl, transferred to a tube with filter (Grace Davison Discovery Science, Columbia, USA) and centrifuged. The filtrate was transferred to a LC-MS/MS vial for

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analysis. Samples were analyzed on an Agilent 1260 LC and an Agilent G6460A QQQ. Samples were separated on an Agilent Zorbax Eclipse XDB-C18 (Santa Clara, CA, USA) 4.6 × 150 mm, 5 μm column by Millipore water with 0.1% heptafluorobutyric acid (v/v, eluent A), and acetonitrile with 0.1% heptafluorobutyric acid (v/v). Flow rate was 0.4 mL min−1, injection volume 5 μL, and column temperature 20 °C. The following gradient was applied: 0 min 5% B, 2 min 10% B, 3 min 20% B, 6 min 20% B, 10 min 50% B, 18 min 90% B with a 9-min postrun at 5% B and linear increases in B between the time steps. Compound detection and quantification was performed as in [57], omitting the solid phase extraction step did not affect recovery. Samples were analyzed for four STX variants (STX, neoSTX, decarbamoylSTX, and decarbamoylneoSTX) and six gonyautoxins (gonyautoxin 1–4, decarbomoyl gonyautoxin 2–3). Calibration standards for all analyzed toxins were obtained from the National Research Council (Canada). 2.5.5. Determination of β-methylamino-L-alanine and 2,4-diaminobutyric acid Lyophilized cyanobacterial material was analyzed for free, soluble bound and total BMAA and DAB as described in [24]. Lypohilized material was extracted in 0.1 M trichloroacetic acid to release free BMAA and DAB, while a part of this extract was dried down, and hydrolysed overnight in 6 M HCl at 105 °C to obtain the soluble bound BMAA and DAB fraction. Another portion of lyophilized material was directly hydrolysed to obtain the total BMAA and DAB contents. Samples were analyzed by LC-MS/MS without derivatisation as described in [18]. 2.6. Qualitative analysis of toxin genes The culture volume of approx. 15 mL was filtered through a 0.45 μm pore diameter nitrocellulose membrane filter (Millipore, USA). Filters with cyanobacterial material were placed into the lysis buffer containing 40 mM EDTA, 400 mM NaCl, 0.75 M sucrose, and 50 mM TRIS-HCl (pH 8.3) and frozen as soon as possible. Genetic material was extracted from the filters according to Giovannoni et al. [27] with some modifications, which improved the extracted DNA quality and quantity, as described by Mankiewicz-Boczek et al. [41]. For the centrifugation, a speed of 13,000 × g instead of 10,000 ×g was used. For the enzymatic lysis step, a final concentration of proteinase K (Fermentas, Lithuania) of 275 μg mL−1 instead of 160 μg mL−1 was used. During the phenol/ chloroform step, a volume of chloroform/isoamyl alcohol (24:1) equal to the volume of supernatant was used. Extracted DNA was adopted as the template for qualitative (PCR, polymerase chain reaction) determination of: cyrA (1105–1179 bp) and cyrJ (578 bp) gene fragments universal for CYN-producing cyanobacteria, anaF (467 bp) a gene fragment universal for ATX-producing cyanobacteria and mcyE (405 bp) a gene fragment universal for MC-producing cyanobacteria. Characteristics of these genes and the primers used for their amplification are included in Table 1. The conditions of PCR amplification of the cyrA gene were optimized according to Kellmann et al. [35]. The PCR was performed in a 20 μL reaction mix containing 1× PCR buffer (Qiagen, Germany), 2.5 mM MgCl2, 0.2 mM dNTPs (Qiagen, Germany), 0.5 μM of each primer, 0.1 mg mL−1

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of BSA, 1 μL of DNA and 0.2 U of Taq DNA polymerase (Qiagen, Germany). The PCR amplification conditions of the cyrJ gene was conducted as described previously by Mankiewicz-Boczek et al. [42]. The PCR was performed in a 20-μL reaction mix containing 1 × PCR buffer, 2.5 mM MgCl2, 0.2 mM dNTPs, 1 μM of each primer (modification in the present study), 1 μL of DNA and 0.2 U (modification in the present study) of Taq DNA polymerase. For both cyrA and cyrJ the initial denaturation step at 94 °C for 3 min was followed by 30 cycles of DNA denaturation at 94 °C for 10 s, primer annealing at 57 °C for 20 s, strand extension at 72 °C for 1 min and final extension step at 72 °C for 7 min. Positive control for PCR analyses of cyrA and cyrJ genes, C. raciborskii strain CS505, was purchased from CSIRO Microalgae Supply Service of the Australian National Algae Culture Collection (ANACC). The PCR amplification conditions of the mcyE gene were similar to those described previously by Mankiewicz-Boczek et al. [43]. The PCR was performed in a 20 μL reaction mix containing 1 × PCR buffer, 3 mM MgCl2, 0.25 mM dNTPs, 0.25 μM of each primer, 0.1 mg mL−1 of BSA, 1 μL of DNA and 1 U of Taq DNA polymerase. The initial denaturation step at 95 °C for 5 min was followed by 30 cycles of DNA denaturation at 94 °C for 30 s, primer annealing at 59 °C for 30 s, strand extension at 72 °C for 1 min and final extension step at 72 °C for 10 min. Positive control for PCR analyses of mcyE gene, M. aeruginosa strain PCC 7806, was purchased from Pasteur Culture Collection (Institut Pasteur, France). The PCR amplification conditions of the anaF gene were similar to those described by Ballot et al. [6,7] with minor modifications. The PCR was performed in a 20 μL reaction mix containing 1× PCR buffer, 2.5 mM MgCl2, 0.2 mM dNTPs, 0.5 μM of each primer, 0.1 mg mL−1 of BSA, 1 μL of DNA, and 1 U of Taq DNA polymerase. The initial denaturation step at 94 °C for 4 min was followed by 30 cycles of DNA denaturation at 94 °C for 10 s, primer annealing at 60 °C (modification in present study) for 20 s, strand extension at 72 °C for 1 min and final extension step at 72 °C for 3 min. Positive control for PCR analyses of anaF gene, the DNA of Cuspidothrix issatschenkoi NIVA CYA 711, was delivered by courtesy of Dr. rer. nat. Andreas Ballot from Norwegian Institute for Water Research. The PCR amplifications were performed with the use of the Eppendorf MasterCycler Gradient thermocycler. The products obtained through PCR amplification were separated by horizontal agarose gel electrophoresis (1.5% agarose, 0.4 μg mL−1 EtBr).

3. Results and discussion 3.1. Molecular identification of species Sequence analysis of 16S rRNA revealed a homology (99–100%) between three studied strains (LBY-Cr, LBO-Cr and LKM-Cr) and C. raciborskii IFCC-CR01 from Turkey (accession no. KY077263). A. gracile LBY-Ag and LBN-Ag revealed homology (100%) with A. gracile HSSASE16 from Egypt (accession no. KT277799). In turn, the 16S rRNA of the LWI-Ag strain showed 98% homology with the strain A. gracile NIVA-CYA 851 from Norway (accession no. LT549450; [4]).

Table 1 Molecular markers and primer sequences used in the present study. Organism

Targeting gene

Primers

Sequence (5′ to 3′)

Length [bp]

Reference

Annealing temperature [°C]

CYN positive cyanobacteria

cyrA

CYLAT-F CYLAT-R cynsulfF cylnamR mcyE-R1 mcyE-S1 atxoaf atxar

ATTGTAAATAGCTGGAATGAGTGG TTAGGGAAGTAATCTTCACAG ACTTCTCTCCTTTCCCTATC GAGTGAAAATGCGTAGAACTTG ATAGGATGTTTAGAGAGAATTTTTTCCC GGGACGAAAAGATAATCAAGTTAAGG TCGGAAGCGCGATCGCAAATCG GCTTCCTGAGAAGGTCCGCTAG

1105–1179

[35]

57

578

[48]

57

405

[43]

59

467

[6,7]

60

cyrJ MC positive cyanobacteria

mcyE

ATX positive cyanobacteria

anaF

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Fig. 1. The intracellular ROS concentrations in human neutrophils (measured by means of DCFDA fluorescence) exposed for 1 h to extracts of C. raciborskii strains: LBY-Cr, LBO-Cr and LKM-Cr (A) and A. gracile strains: LBY-Ag, LBN-Ag and LWI-Ag (B) isolated in Poland expressed as percentage of control. Bars represent mean ± SD from five independent experiments corresponding to different donors. Asterisks represent statistically significant difference to the control (* - p b 0.05; ** - p b 0.01; Wilcoxon signed-rank test).

3.2. Morphological characteristics No intraspecific morphological differences between particular strains of C. raciborskii and A. gracile investigated in the present study were observed (Supplementary Fig. S1). Trichomes of both species were solitary, straight, cylindrical and slightly to distinctly constricted at the cross-walls with cells cylindrical to barrel-shaped, longer than wide. Terminal cells of A. gracile were often elongated and narrowed; heterocytes were common, located intercalary and solitary. Several heterocytes were observed in the case of long trichomes. Short, cylindrical, solitary akinets with a characteristic cup-shaped sheath formation were seen only rarely under culture conditions. Apical cells of C. raciborskii were distinctly narrowed. Terminal heterocytes were commonly observed at one or both ends of trichomes with a characteristic elongated drop-lake shape. Akinets were rare, oval, located near the ends of trichomes usually adjacent to the heterocytes.

3.3. In vitro toxicity As demonstrated by the present study, the investigated cell-free cyanobacterial extracts exerted distinctively different effects on human neutrophils. Compared to the control, cells exposed for 1 h to extracts of C. raciborskii LBY-Cr, A. gracile LBN-Ag and A. gracile LWI-Ag exhibited a steady increase in ROS level which had exceeded over 25%, 11.1 and 8.1% by the end of monitored period, respectively (Fig. 1). By using this assay we were able to evaluate which extract can induce oxidative stress, an imbalance between ROS generation and biological system ability to readily detoxify the reactive intermediates or to repair the resulting damage [29]. This phenomenon has already been shown to be triggered in animal cells by the vast majority of cyanotoxins including CYN [55], GAA [50], STX [47], ATX [60] and BMAA [40]. The potential detrimental outcomes of oxidative stress include DNA damage, protein modifications, lipid peroxidation, and necrotic or apoptotic cell death [29]. The present study found a significant increase of lipid peroxidation, as measured by means of malondialdehyde (MDA) formation, for neutrophils exposed to extracts of C. raciborskii LBY-Cr and A. gracile LBN-Ag (Fig. 2). Peroxidation of lipids is a chain reaction initiated by hydrogen abstraction or addition of an oxygen radical, resulting in the oxidative damage of polyunsaturated fatty acids. If not terminated fast enough a decrease in membrane fluidity and in the barrier functions of membranes is induced, final products of peroxidation (predominantly MDA and 4-hydroxy-2-nonenal) can induce genotoxicity, and cell death is promoted [3,12]. As demonstrated, extracts of C. raciborskii LBY-Cr and A. gracile LBN-Ag also affected the viability of human neutrophils; following 1 h of exposure - the OD570 of purple formazan generated by active mitochondrial dehydrogenases was decreased by as much as 23.6 and 15.6%, respectively (Fig. 4). Importantly, extracts obtained from other

strains of C. raciborskii (LBO-Cr and LKM-Cr) and A. gracile LBY-Ag had no effect on intracellular ROS levels (Fig. 1), lipid peroxidation (Fig. 2) and cell viability (Fig. 3). Although the presence of exudates of A. gracile LWI-Ag caused a slight increase in ROS level, other effects were insignificant indicating that neutrophils successfully coped with the redox imbalance (Figs. 1–3). The findings of the present study are in line with previous observations that C. raciborskii LBY-Cr contain unidentified metabolite(s) exhibiting pro-necrotic and pro-apoptotic action in human cells [54,55], and further elucidate that these effects are likely to be a result of an intracellular redox imbalance. They also indicate that the toxicity of C. raciborskii and A. gracile in Polish freshwaters is strain-specific. Similarly, Antal et al. and Acs et al. [1,2] also observed that the magnitude of in vitro adverse effects exerted by C. raciborskii isolated from Lake Balaton (Hungary) was highly strain-specific [1,2]. If one considers the short-term exposure model applied in our study, the magnitude of the observed effects highlights the bioactive power of C. raciborskii LBY-Cr and A. gracile LBN-Ag exudates. The in vitro assays performed in the present investigation indicated that metabolite(s) produced by one strain of C. raciborskii (LBY-Cr) and one strain of A. gracile (LBN-Ag) are human cell membrane permeable, increase intracellular levels of ROS above the adaptive responses causing oxidative damage, specifically peroxidation of lipids, and can eventually lead to cell death. These findings strongly advocated a need to screen the investigated strains for the production of different cyanobacterial toxins.

Fig. 2. The level of lipid peroxidation (measured by means of MDA concentration) in human neutrophils exposed for 1 h to extracts of C. raciborskii (black bars) and A. gracile (grey bars) strains isolated in Poland expressed as percentage of control. Bars represent mean ± SD from five independent experiments corresponding to different donors. Asterisks represent statistically significant difference to the control (* - p b 0.05; ** - p b 0.01; Wilcoxon signed-rank test).

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Fig. 3. The viability of human neutrophils exposed for 1 h to extracts of C. raciborskii (black bars) and A. gracile (grey bars) strains isolated in Poland measured by means of mitochondrial activity in MTT assay generating the formazan crystals absorbing at 570 nm, and expressed as percentage of control. Bars represent mean ± SD from five independent experiments corresponding to different donors. Asterisk represents statistically significant difference to the control (p b 0.05; Wilcoxon signed-rank test).

3.4. Production of cylindrospermopsin and guanidinoacetate As found, none of the investigated C. raciborskii and A. gracile strains produced CYN nor detectable levels of GAA, and did not possess either cyrA or cyrJ genes (Fig. 4). The former gene encodes an amidinotransferase responsible for transferring a guanidino group from

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arginine onto glycine. This reaction, a first step in the formation of the carbon skeleton of CYN, consequently results in the production of GAA [35, 48]. The cyrJ gene, in turn, regulates the sulfation at C-12 essential to complete CYN biosynthesis [48], and its screening has been previously demonstrated as a convenient method to confirm or exclude CYN production in cyanobacteria [42,44]. The present study indicates that CYN biosynthesis is not even initiated in the tested strains, and that the observed in vitro toxicity cannot be explained by potential CYN intermediates as they are not produced. It still remains unclear whether the tested strains of C. raciborskii (as well as other European strains) never acquired the ability to produce CYN or lost it over time under certain environmental conditions. It has been phylogenetically supported that Europe was likely to have been the last continent to be colonized by this species, after its American origin and subsequent spread into Africa, Asia and Australia, and suggested that this dispersal route may affect the distribution of toxin production [49]. However, further studies are necessary to understand the exact events that influenced the observed distribution of CYN genes in C. raciborskii. Nevertheless, despite the Europe-wide distribution of C. raciborskii and extensive research conducted over the years [53,62], to date no CYN-producing strain has been identified on this continent [62]. Although one report associated CYN occurrence in Lake Aleksandrovac in Serbia with a C. raciborskii dominated bloom, investigations were not performed on the isolated strain [17] while another study found C. raciborskii from the same reservoir to be toxic but CYN-negative [67]. Therefore, it remains unclear whether any Serbian strain of this species is capable of CYN biosynthesis. A. gracile, in turn, has already been identified as a producer of CYN in Europe but this finding is so far limited to certain freshwater lakes in Poland [36]. In the present study the LBY-Ag strain isolated originally from Lake Bytyńskie did not produce CYN although the water from this reservoir was previously shown to contain detectable levels of the toxin and the cyrJ gene [36]. It is possible, however, that toxin producers may co-

Fig. 4. The results of qualitative determination of: cyrA (1105–1179 bp) and cyrJ (578 bp) gene fragments universal for CYN-producing cyanobacteria, anaF (467 bp) gene fragment universal for ATX-producing cyanobacteria and mcyE (405 bp) gene fragment universal for MC-producing cyanobacteria in C. raciborskii (-Cr) and A. gracile (-Ag) strains isolated in Poland; Control “+”: isolate of strains C. raciborskii CS505 (for cyrA and cyrJ genes), Cuspidothrix issatschenkoi NIVA CYA 711 (for anaF gene) and M. aeruginosa PCC 7806 (for mcyE gene); Control “−”: sterile Millipore water; M – DNA marker M100-500; M100-700; M100-3000 (DNA Gdańsk, Poland).

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occur with toxin-negative strains; this has also been evidenced for STX in A. gracile [7]. The lack of GAA production in the strains tested in the present study further indicates that this compound may not be a general cyanobacterial metabolite, and that only certain non-CYN producing strains may synthesize it at relatively low concentrations [8].

3.5. Production of microcystins The cyanobacterial strains investigated in the present study did not produce any MCs, as confirmed by the lack of any analytical detection of its three most common analogues (MC-LR, MC-YR and MC-RR), and by no presence of mcyE gene (Fig. 4). These toxins have never been confirmed for species classified within the former Aphanizomenon genus (now under revision; for details see e.g. [14]) with the exception of the Moroccan strain of Sphaerospermopsis (Aphanizomenon) aphanizomenoides for which MC production was presumed by means of analytical determination [65]. Their biosynthesis has also never been found in C. raciborskii. A single study demonstrated that a strain isolated from the Bir M'cherga reservoir in Tunisia possessed two segments from the mcy gene cluster (mcyA and mcyE genes) although the toxin was not analytically detected [26]. This finding created a necessity to screen other strains, particularly those revealing toxicity but not producing any anticipated cyanotoxin, for the presence of MC on both the molecular and analytical level.

3.6. Production of anatoxin-a and saxitoxins As found, none of the isolated C. raciborskii and A. gracile strains produced detectable concentrations of ATX nor did they contain the anaF gene (Fig. 4). To date, the production of ATX has never been confirmed for any A. gracile and C. raciborskii strain although this issue has been very rarely addressed [14,62,68]. Recent studies, however, have demonstrated that extracts of C. raciborskii strains isolated from Lake Balaton in Hungary, analytically confirmed not to contain ATX, produce unknown neurotoxic metabolites that evoke cholinergic inhibitory effects similar to those of ATX [1,69,70]. These findings highlighted the need to conduct further screening on ATX presence in strains occurring in different European locations. The assessment of ATX production in strains tested in the present study was based on analytical investigations coupled with PCR targeting of the anaF gene tailoring a crucial Michael type cyclization in ATX biosynthesis [45,46]. This approach was necessary as ATX can be easily transformed to its dihydro derivatives, and underestimated or not detected at all [59]. Overall, the present study investigated the presence of ten different STX analogues - GTX1, dcGTX2, GTX2, dcGTX3, GTX3, GTX4, dcNEO, NEO, dcSTX and STX. Similarly to ATX, none were found for any A. gracile or C. raciborskii strain. To our knowledge, this is the first time the production of STXs has been analyzed in cyanobacteria isolated in Poland. Both, C. raciborskii and A. gracile are known to be potential STXs producers; the ability of their biosynthesis is, however, geographically diversified. In the case of C. raciborskii, STXs were only confirmed for strains associated with South America [13,30,31]. However, a recent study reported the presence of STX in bloom dominated by C. raciborskii in Greece, but no final conclusion can be drawn without molecular and analytical investigations on isolated strains [28]. Studies based on such a dualistic approach have already excluded STX biosynthesis in European C. raciborskii strains from various locations, although none have tested those occurring in Poland [1,15]. In fact, as far as the Central European region is concerned, only Hungarian C. raciborskii strains were investigated [1]. In turn, STX-producing strains of A. gracile have been identified in Europe, in German, French and Spanish freshwaters [6,7,15,38]; one STX-positive strain (misclassified originally as Aphanizomenon flos-aquae) has also been isolated from the US [14].

3.7. Production of β-methylamino-L-alanine and 2,4-diaminobutyric acid None of the investigated strains of C. raciborskii and A. gracile were found to produce detectable levels of this toxin. Although BMAA has been identified in European freshwaters [22], information on potential producers is still scarce. To our knowledge this is the first time that BMAA has been analyzed for cyanobacteria associated with the Central European region, further highlighting the need for further monitoring for the presence of this compound. The research on BMAA is highly important due to the neurodegenerative potency of this non-essential amino acid and experimentally-evidenced adverse effects exerted on aquatic animals [19]. It was originally found that Nostoc sp. symbiotic to cycads are responsible for BMAA production, and the dietary intake of this compound was first suggested as an aetiological factor in amyotrophic lateral sclerosis-Parkinsonism dementia complex among the Chamorro people on Guam [16]. Later, it was hypothesized to play a potential role in the global epidemiology of neurodegenerative diseases [11]. Some studies confirmed the presence of BMAA in certain cyanobacteria whereas others did not, the discrepancies are linked to invalid methodology used in BMAA quantification [20,21,23]. It is still unclear to which extent cyanobacteria contribute to BMAA occurrence in freshwaters and whether other representatives of phytoplankton (e.g. diatoms) may be more important producers of this amino acid [23,34,58]. In the present study, the most accepted and validated method for BMAA identification (LC MS/MS) in biological samples was employed [18,20]. None of investigated strains of C. raciborskii contained detectable levels of DAB. The toxin was, however, found for A. gracile LWI-Ag at concentrations below the level of quantification (1.0 μg g−1 dry weight). The exact health risks arising from DAB occurrence are not known as its toxicology is not yet fully assessed. Even though an extract containing DAB induced slightly increased intracellular ROS levels it did not increase lipid peroxidation or decrease neutrophil survival (Figs. 1– 3). It is unclear whether environmental conditions, and of what kind, can trigger DAB production, although a single study employing M. aeruginosa demonstrated that intracellular DAB content increases over the nutrient concentration [73]. Further experimental and/or in-field studies would be necessary to test whether such a phenomenon also occurs in A. gracile and whether eutrophication can promote the production of this compound. Similarly to BMAA, the exact pathway of biosynthesis for DAB is unknown and requires characterization on genetic level. Nevertheless, this is the first time DAB that has been identified for cyanobacteria occurring in Polish freshwater. Although different cells of cyanobacteria of European origin, e.g. M. aeruginosa or Nodularia spumigena have already been confirmed to contain this compound [37], the present study is the first to show that A. gracile can be its potent producer in this geographical area. Wider-scale investigations are necessary to determine how frequently this species contributes to DAB occurrence in European surface waters. 4. Conclusions In summary, the present study assessed the toxicities of C. raciborskii and A. gracile isolated from freshwater lakes of Western Poland. As demonstrated, exudates of one C. raciborskii and one A. gracile strain revealed significant cytotoxic action in human neutrophils through induction of oxidative stress. Although these findings implicate the production of certain powerful toxic compounds by these cyanobacteria, cyanotoxin screening yielded mostly negative results. None of the tested strains produced GAA, CYN, ATX, STXs, MCs nor BMAA. The production of low concentrations of DAB was confirmed only for one A. gracile strain but its extract did not induce lipid peroxidation or cell death. This study indicates the production of as yet unknown metabolite(s) that may pose a relevant threat to human health by strains of C. raciborskii and A. gracile occurring in Polish lakes, and adds to the general understanding of toxicity of European strains of both species.

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Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.algal.2017.02.011.

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Contributions

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Piotr Rzymski ([email protected]) had contributed to the conception and design of the study, acquisition of data, drafting the article and final approval of the version to be submitted, and takes responsibility for the integrity of the work as a whole; Barbara Poniedziałek, Joanna Mankiewicz-Boczek, Elisabeth J. Faassen, Tomasz Jurczak, Ilona Gągała-Borowska, Andreas Ballot, Miquel Lürling and Mikołaj Kokociński had contributed to acquisition of data, provision of study material, drafting the article and final approval of the version to be submitted. Acknowledgments Piotr Rzymski is supported by the Foundation for Polish Science (FNP) (START 091.2016). References [1] A. Acs, A.W. Kovács, J.Z. Csepregi, N. Törő, G. Kiss, J. Győri, A. Vehovszky, N. Kováts, A. Farkas, The ecotoxicological evaluation of Cylindrospermopsis raciborskii from Lake Balaton (Hungary) employing a battery of bioassays and chemical screening, Toxicon 70 (2013) 98–106. [2] O. Antal, M. Karisztl-Gácsi, A. Farkas, A. Kovács, A. Acs, N. Töro, G. Kiss, M.L. Saker, J. Gyori, G. Bánfalvi, A. Vehovszky, Screening the toxic potential of Cylindrospermopsis raciborskii strains isolated from Lake Balaton, Hungary, Toxicon 57 (2011) 831–840. [3] A. Ayala, M.F. Muñoz, S. Argüelles, Lipid peroxidation: production, metabolism, and signaling mechanisms of malondialdehyde and 4-hydroxy-2-nonenal, Oxidative Med. Cell. Longev. 2014 (2014) 360438. [4] A. Ballot, L. Cerasino, V. Hostyeva, S. Cirés, Variability in the sxt gene clusters of PSP toxin producing Aphanizomenon gracile strains from Norway, Spain, Germany and North America, PLoS One 11 (2016), e0167552. . [5] A. Ballot, J. Ramm, T. Rundberget, R.N. Kaplan-Levy, O. Hadas, A. Sukenik, C. Wiedner, Occurrence of non-cylindrospermopsin-producing Aphanizomenon ovalisporum and Anabaena bergii in Lake Kinneret (Israel), J. Plankton Res. 33 (2011) 1736–1746. [6] A. Ballot, J. Fastner, M. Lentz, C. Wiedner, First report of anatoxin-a-producing cyanobacterium Aphanizomenon issatschenkoi in northeastern Germany, Toxicon 56 (2010) 964–971. [7] A. Ballot, J. Fastner, C. Wiedner, Paralytic shellfish poisoning toxin-producing cyanobacterium Aphanizomenon gracile in Northeast Germany, Appl. Environ. Microbiol. 76 (2010) 1173–1180. [8] Á. Barón-Sola, S. Sanz-Alférez, F.F. del Campo, First evidence of accumulation in cyanobacteria of guanidinoacetate, a precursor of the toxin cylindrospermopsin, Chemosphere 119 (2015) 1099–1104. [9] Y. Bar-Yosef, A. Sukenik, O. Hadas, Y. Viner-Mozzini, A. Kaplan, Enslavement in the water body by toxic Aphanizomenon ovalisporum, inducing alkaline phosphatase in phytoplanktons, Curr. Biol. 20 (2010) 1557–1561. [10] C. Bernard, M. Harvey, J.F. Briand, R. Biré, S. Krys, J.J. Fontaine, Toxicological comparison of diverse Cylindrospermopsis raciborskii strains: evidence of liver damage caused by a French C. raciborskii strain, Environ. Toxicol. 18 (2003) 176–186. [11] W.G. Bradley, D.C. Mash, Beyond Guam: the cyanobacteria/BMAA hypothesis of the cause of ALS and other neurodegenerative diseases, Amyotroph. Lateral Scler. 10 (Suppl. 2) (2009) 7–20. [12] P.C. Burcham, Genotoxic lipid peroxidation products: their DNA damaging properties and role in formation of endogenous DNA adducts, Mutagenesis 13 (1998) 287–305. [13] R.L. Carneiro, A.B. Pacheco, S.M. de Oliveira e Azevedo, Growth and saxitoxin production by Cylindrospermopsis raciborskii (cyanobacteria) correlate with water hardness, Mar. Drugs 11 (2013) 2949–2963. [14] S. Cirés, A. Ballot, A review of the phylogeny, ecology and toxin production of bloom-forming Aphanizomenon spp. and related species within the Nostocales (cyanobacteria), Harmful Algae 54 (2016) 21–43. [15] S. Cirés, L. Wörmer, A. Ballot, R. Agha, C. Wiedner, D. Velázquez, M.C. Casero, A. Quesada, Phylogeography of cylindrospermopsin and paralytic shellfish toxin-producing nostocales cyanobacteria from Mediterranean Europe (Spain), Appl. Environ. Microbiol. 80 (2014) 1359–7130. [16] P.A. Cox, S.A. Banack, S.J. Murch, Biomagnification of cyanobacterial neurotoxins and neurodegenerative disease among the Chamorro people of Guam, Proc. Natl. Acad. Sci. U. S. A. 100 (2003) 13380–13383. [17] N. Dorđević, S.B. Simić, A.R. Ciric, First identification of the cylindrospermopsin (CYN) producting by cyanobacterium Cylindrospermopsis raciborskii (Woloszynska) Seenayya & Subba Raju in Serbia, Fresenius Environ. Bull. 24 (2015) 3736–3742. [18] E.J. Faassen, M.G. Antoniou, W. Beekman-Lukassen, L. Blahova, E. Chernova, C. Christophoridis, A. Combes, C. Edwards, J. Fastner, J. Harmsen, A. Hiskia, L.L. Ilag, T. Kaloudis, S. Lopicic, M. Lürling, H. Mazur-Marzec, J. Meriluoto, C. Porojan, Y. VinerMozzini, N. Zguna, A collaborative evaluation of LC-MS/MS based methods for

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