Harmful Algae 10 (2011) 668–675

Cylindrospermopsin production and release by the potentially invasivecyanobacterium Aphanizomenon ovalisporum under temperatureand light gradients

Samuel Cires a,1, Lars Wormer a, Jesus Timon a, Claudia Wiedner b,2, Antonio Quesada a,*a Departamento de Biologıa, C/Darwin, 2, Universidad Autonoma de Madrid, ES-28049 Madrid, Spainb Leibniz-Institute of Freshwater Ecology and Inland Fisheries, Department of Limnology of Stratified Lakes, Alte Fischerhutte 2, D-16775 Stechlin, Germany

A R T I C L E I N F O

Article history:

Received 11 April 2011

Received in revised form 16 May 2011

Accepted 16 May 2011

Available online 25 May 2011

Keywords:

Aphanizomenon ovalisporum

Cylindrospermopsin

Invasive

Light intensity

Release

Temperature

A B S T R A C T

The growth rates, production and release of the potent cytotoxin cylindrospermopsin (CYN) were

studied in batch and semi-continuous cultures of Aphanizomenon ovalisporum (Cyanobacteria;

Nostocaceae) strains UAM 289 and UAM 290 from Spain, over a gradient of temperatures (10–40 8C)

and irradiances (15–340 mE m�2 s�1). This species grew in temperatures ranging from 15 8C to 35 8C as

well as under all irradiances assayed. The growth rates ranged from 0.08 d�1 to 0.35 d�1, and the

maximum growth was recorded above 30 8C and at 60 mE m�2 s�1. CYN was produced under all

conditions where net growth occurred. Total CYN reached up to 6.4 mg mg�1 dry weight, 2.4 mg mm�3

biovolume, 190.6 fg cell�1 and 0.5 mg mg�1 chlorophyll a. Although CYN concentrations varied only 1.9-

fold within the 15–30 8C range, a drastic 25-fold decrease was observed at 35 8C. The irradiance induced

up to 4-fold variations, with maximum total CYN measured at 60 mE m�2 s�1. An elevated extracellular

CYN share ranging from 20% to 35% was observed during the exponential growth phase in most

experiments, with extreme temperatures (15 and 35 8C) being related to the highest release (63% and

58%, respectively) and without remarkable influence of irradiance. Growth did not have a direct

influence on either CYN production or release throughout the entire range of experimental conditions.

Our study demonstrates a strong and stable production and release of CYN by A. ovalisporum along field-

realistic gradients of temperature and light, thus becoming a predictive tool useful for the management

of water bodies potentially affected by this ecologically plastic cyanobacterium.

� 2011 Elsevier B.V. All rights reserved.

Contents lists available at ScienceDirect

Harmful Algae

jo u rn al h om epag e: ww w.els evier .c o m/lo cat e/ha l

1. Introduction

Under certain environmental conditions, planktonic cyanobac-teria may develop massive proliferations (‘‘blooms’’) (Mur et al.,1999). This worldwide phenomenon poses a public health riskbecause of the ability of some cyanobacterial strains to producecyanotoxins, which are diverse secondary metabolites responsiblefor severe human intoxications (Codd et al., 2005).

Among these cyanotoxins, the alkaloid cylindrospermopsin(CYN) is of increasing concern due to its potent cytotoxic effectsand the growing number of worldwide locations in which CYN hasbeen reported during the last years. First described from AustralianCylindrospermopsis raciborskii (Ohtani et al., 1992), CYN inhibits

* Corresponding author. Tel.: +34 914978181; fax: +34 914978344.

E-mail addresses: samuel.cires@uam.es (S. Cires), antonio.quesada@uam.es

(A. Quesada).1 Tel.: +34 914978177.2 Present address: Brandenburg University of Technology, Cottbus, Department

of Freshwater Conservation, Seestr. 45, D-15526 Bad Saarow, Germany.

1568-9883/$ – see front matter � 2011 Elsevier B.V. All rights reserved.

doi:10.1016/j.hal.2011.05.002

glutathione, protein synthesis and cytochrome P450 resulting inhepatotoxic, neurotoxic and general cytotoxic effects in mammals(Pearson et al., 2010), as well as potentially interfering with DNAstructure (Shaw et al., 2000). CYN poisoning was responsible forthe Palm Island Mystery Disease (Griffiths and Saker, 2003), asevere hepatitis-like illness affecting 148 people that occurred in1979 on Palm Island (Queensland, Australia). Further, the toxicityof CYN-containing cyanobacterial extracts has been demonstratedin aquatic organisms including zooplankton (Nogueira et al., 2004),mollusks (Kinnear et al., 2007a) and amphibians (Kinnear et al.,2007b).

So far, CYN has been reported in freshwaters worldwide,including Oceania (Shaw et al., 1999), America (Burns et al., 2002),Asia (Li et al., 2001a) and Europe (Quesada et al., 2006), with anincreasing number of reports in the European freshwaters in thelast two years (Blahova et al., 2009; Brient et al., 2009; Kokocinskiet al., 2009). In most waterbodies, the presence of CYN ischaracterised by a high dissolved fraction (from 6% to 100%) oftotal CYN (Wormer et al., 2009). In fact, and unlike the morestudied microcystins (MCYST), dissolved CYN tends to accumulate

S. Cires et al. / Harmful Algae 10 (2011) 668–675 669

in the field because of elevated extracellular release by CYN-producing cyanobacteria (Hawkins et al., 2001; Preussel et al.,2009; Mazmouz et al., 2010) together with limited photodegrada-tion (Wormer et al., 2010) and biodegradation (Wormer et al.,2008).

Besides C. raciborskii, CYN is produced by a number offilamentous cyanobacteria (Pearson et al., 2010). For each of theCYN-producing species, both CYN-producing and non-producingstrains have been found, the exception being the freshwaterplanktonic Nostocacean Aphanizomenon ovalisporum, of which allthe strains reported to date synthesize CYN regardless of theirgeographical origin (Stuken et al., 2009).

A. ovalisporum Forti (1911) was first described from lakeKutchuk Tchekmedje near Istanbul (Turkey), and was laterreported in subtropical areas of Australia (Shaw et al., 1999) andFlorida in the USA (Yilmaz et al., 2008), as well as in theMediterranean areas of the Middle East (Banker et al., 1997) andEurope (Gkelis et al., 2005; Quesada et al., 2006). Dense blooms ofthis species have been reported in Lake Kinneret (Israel) (Bankeret al., 1997), in newly constructed lakes from Queensland(Australia) (Shaw et al., 1999), in the Arcos reservoir (Spain)(Quesada et al., 2006), and in the volcanic Lake Albano (Italy)(Messineo et al., 2010). In Europe, these blooms were concurrentwith 9.4–18 mg CYN L�1 (Messineo et al., 2010; Quesada et al.,2006), and up to 120 mg CYN L�1 (Shaw et al., 1999) in Australia,thus posing a serious health risk for recreational water usage andconsumption.

Concerning the ecophysiology of A. ovalisporum, strains fromthe Israeli Lake Kinneret showed a preference for low to moderatelight irradiances (Hadas et al., 2002) and temperature optima of26–30 8C. More recently, Mehnert et al. (2010) reported anoptimum of 32.8 8C for the Spanish strain UAM 290 (used in ourstudy), modelling a potential northwards spreading of this speciesunder the global warming scenario.

The regulation of CYN production and release by environmentalfactors was first studied for tropical C. raciborskii, and morerecently for A. ovalisporum and Aphanizomenon flos-aquae. CYNproduction was influenced by temperature in A. flos-aquae

(Preussel et al., 2009) and C. raciborskii (Saker and Griffiths,2000), and by light intensity in C. raciborskii (Dyble et al., 2006).Regarding extracellular CYN release, only few studies are availableconfirming dissolved-CYN concentrations above 50% of the totalCYN in exponentially growing semi-continuous cultures of A. flos-

aquae (Preussel et al., 2009) and in C. raciborskii during thestationary phase (Hawkins et al., 2001; Saker and Griffiths, 2000).The release of CYN in A. flos-aquae was suggested to be stress-induced (Preussel et al., 2009).

Particularly in A. ovalisporum, shifts in the CYN cellular contenthave been studied in detail in relation to sulphate and phosphorus(Bacsi et al., 2006). Although the transcription levels of the genesaoaA and aoaC – involved in CYN synthesis in A. ovalisporum – wereestimated in relation to light intensity and nitrogen availability(Shalev-Malul et al., 2008), experiments monitoring the quantita-tive variations of intracellular CYN content and CYN release in thisspecies under light and temperature gradients are still lacking.Furthermore, given that two strains of a single CYN-producingspecies might show different responses on temperature and light-dependent CYN production (Preussel et al., 2009), studies like thepresent including at least two strains are still necessary.

In this context, we characterised the growth response, and CYNproduction and release in batch and semi-continuous cultures oftwo A. ovalisporum strains (UAM 289 and UAM 290) from Spanishfreshwaters in relation to shifts in temperature and light intensity.The aims of this study are (1) to achieve a better understanding ofthe ecophysiology of the potentially invasive species A. ovalis-

porum; (2) to determine how realistic conditions of temperature

and light intensity affect CYN dynamics in A. ovalisporum; and (3)to find a global relationship between CYN production/release andgrowth rates.

2. Materials and methods

2.1. Culture experiments

A. ovalisporum strains UAM 289 and UAM 290 were isolatedfrom a pond located inside the urban park Juan Carlos I (Madrid,Spain) as described in Wormer et al. (2008). The cultures weresubsequently kept in BG110-medium (Rippka et al., 1979) at aconstant temperature of 28 8C under continuous white lightillumination of 60 mE m�2 s�1 until they were used for light andtemperature experiments. These two strains showed a 100%similarity for cpcBA-IGS (NCBI GenBank ID: HQ259628 for UAM289, obtained in this study following Ballot et al., 2010; GenBankID: EF529468 for UAM 290, obtained by Stuken et al., 2009) with analignment length of 593 bp, and for the 16S rRNA gene (GenBankID: HQ259629 for UAM 289, obtained in this study following Ballotet al., 2008; GenBank ID: EF529489 for UAM 290, obtained byStuken et al., 2009) with an alignment length of 1225 bp.

The light-intensity experiments were performed under a batchregime for four weeks. For this, the strain UAM 289 was grown intriplicate in 2-L Erlenmeyer flasks containing 1 L of BG110-mediumat a constant temperature of 28 8C under continuous white lightillumination provided by cool fluorescent lights (Phillips TLD 36W/33) and supplemented with sterile air bubbling. Four lightintensities of 15, 60, 200 and 340 mE m�2 s�1 were used in theexperiments. Before beginning each experiment, the cultures wereacclimated to the experimental conditions for at least threegeneration-times.

The temperature experiments included batch-grown and semi-continuous cultures to ensure that our results are comparable withpreviously reported work on CYN production independent of theculturing regime used in those studies. For both cultures, aliquotsof strain UAM 290 were grown in triplicate in 250-mL Erlenmeyerflasks containing 100 mL of Z8-medium (Kotai, 1972) supplemen-ted with 164 nM biotin, 30 nM cyanocobalamine and 119 nMthiamine hydrochloride, which were kept inside growth chambers(BINDER, KBW 400, Germany) with dimmable illumination(OSRAM, Lumilux 18 W/865XT Cool Daylight, Germany) simulat-ing 12:12 h light:dark photoperiods at a light intensity of80 mE m�2 s�1 and continuously shaken at 80–100 r.p.m.

The semi-continuous temperature experiments were carriedout according to the turbidostat principle (Mehnert et al., 2010), bydiluting cultures every second day to an optical density (O.D.750nm)of 0.1. Temperatures from 10 to 40 8C were assayed at intervals of5 8C. Additionally, four-week batch temperature experiments werecarried out at 15, 20, 27.5 and 30 8C, being started with dilutedaliquots of semi-continuous cultures that were previouslyacclimated to each of the temperatures.

Two additional semi-continuous experiments were performedat 30 8C and under 80 mE m�2 s�1. The first experiment examinedthe culture-medium-dependent CYN content by growing UAM 290with BG110-medium, whereas the second experiment examinedthe strain-dependent CYN content by growing UAM 289 withmodified Z8-medium. Again, before the onset of each experiment,cultures were acclimated to the experimental conditions until thegrowth rates were stable.

2.2. Growth parameters

During the experimental period, samples were periodicallytaken for the determination of growth parameters (the opticaldensity at 750 nm, dry weight, biovolume and chlorophyll a

S. Cires et al. / Harmful Algae 10 (2011) 668–675670

concentration). Samples from the batch experiments (light andtemperature experiments) were taken every third day during the4-week experimental period, whereas in the semi-continuousexperiments, samples were taken from four consecutive dilutiondays. Afterwards, samples were processed as follows:

Optical density at 750 nm (O.D.750nm) was measured using aUV/VIS spectrophotometer (Lambda 2, Perkin Elmer, USA).

The dry weight (DW) in the light-intensity experiments wasdetermined from pre-desiccated glass microfibre filters GF/F(Whatman, England), saturated with culture material (5–25 mL)after low-vacuum filtration. The filters were desiccated at 40 8Cand periodically weighed until reaching a constant weight(typically in 24 h). A linear relationship between O.D.750nm andDW was calculated for the light intensity experiments(y = 0.4712x; R2 = 0.95; p < 0.05), and was used to estimate theDW from the O.D.750nm values obtained in all the experiments.

The chlorophyll a (Chl a) concentration was determined frombiomass-saturated GF/F filters (containing 5–25 mL of culturematerial) that were kept at �20 8C until analysis. The filters wereextracted twice with 90% (v/v) methanol with the aid of anultrasonic bath Ultrasonics (J.P. Selecta, Spain). The extract wassubsequently recovered by centrifugation (1385 � g, 15 min) in aLabfuge AE centrifuge (Heraeus Sepatech GmbH, Germany). TheO.D.750nm and O.D.665nm of the extract were measured by a Multi-Spec 1501 UV/VIS spectrophotometer (Shimadzu, Japan) andconverted into Chl a concentrations according to Marker et al.(1980).

The cell concentrations and biovolumes were determined in thetemperature experiments from samples previously fixed with acidlugol, and sedimented following the Utermohls procedure(Utermohl, 1958). The cells were counted under an invertedmicroscope Leica DM IL (Leica Microsystems, Germany) at 400�magnification until reaching statistically significant numbers ofcounting units. Average cell biovolumes were estimated byassimilating cells to regular geometric bodies, these beingmultiplied by cell concentrations thus obtaining the cyanobacter-ial biovolume (mm3) per volume of culture medium (L). A linearrelationship between the O.D.750nm and the cyanobacterialbiovolume could be established (y = 1345.6x; R2 = 0.94; p < 0.05)by including the samples from all of the temperatures (n = 34), theequation being used to estimate the biovolumes from O.D.750nm

values in all the experiments.The maximum specific growth rate was estimated using the

O.D.750nm values obtained for each experimental condition. Thegrowth rate in semi-continuous experiments was determinedaccording to the following equation:

m ¼ lnXDt

X0

� �� 1

Dt; (1)

where XDt is the biomass (O.D.750nm) after cultivation time t, X0 isbiomass after dilution and Dt is the time between two dilutions.The mean values and standard deviation were calculated from theO.D.750nm obtained from four consecutive dilution dates.

For the batch experiments (light-intensity and temperature),the growth curves were adjusted to a 4-parameters logistic model(p < 0.05), except for the 15 8C-experiment in which the bestfitting was achieved by a quadratic model (p < 0.05). Themaximum growth rates were calculated by applying Eq. (1) tothe modelled data in the exponential-growth phase of each of thecurves. Finally, 95% confidence intervals for the logistic adjustmentwere considered to estimate the same intervals for the maximumgrowth rate of each experiment. None of the batch culturesreached the stationary phase during the 4-week period (data notshown), therefore all data presented hereafter refer to theexponential growth phase.

2.3. Cylindrospermopsin quantification

CYN was monitored in cultures according to the same samplingapproach used for the growth parameters (see Section 2.2). TheCYN concentrations in sestonic and dissolved fractions wereanalysed separately in all experiments. The term ‘‘total CYN’’,hereafter, refers to the sum of both fractions.

The CYN in the sestonic fraction was determined from thebiomass saturated GF/F filters (containing 5–25 mL of culturematerial) after low-vacuum filtration. The CYN retained in filtersfrom light-intensity experiments was extracted according toTorokne et al. (2004). The sestonic CYN from the temperatureexperiments was extracted using a similar procedure, although theextractant in this case was Milli-Q water that was suitable foranalysis by mass-spectrometry.

The dissolved CYN was determined from the filtrate obtainedafter the low-vacuum filtration. In the case of the light-intensityexperiments, the samples were pre-concentrated by solid-phaseextraction according to Wormer et al. (2009), and the elutedsolution was filtered through 0.45 mm pore-size nylon filters(Teknokroma, Spain) and placed in chromatography vials. For thetemperature experiments, the filtrate was directly filtered through0.45 mm pore-size nylon filters and placed in chromatographyvials without any other preparation.

The CYN quantification in the light-intensity experiments (bothsestonic and dissolved) was performed on a Waters Alliance 2695HPLC system with a 996 PDA detector equipped with a WatersSpherisorb 5-mm ODS2 column according to the protocol describedby Torokne et al. (2004). The CYN from the temperature experimentswas quantified by LC–MS/MS on a Varian 500 MS Ion Trap MassSpectrometer. The chromatographic separation was based on themethod proposed by Ballot et al. (2010), modified by using a 3-mm,150 mm � 2 mm Pursuit 3 C18 column (Varian, USA) with water (A)and methanol (B) as mobile phases, both containing 5.0 mMammonium formate and 0.2% formic acid. The gradient programwas run using the following parameters: start with 5% B, 5% to 30% Bover 10 s, 30% to 70% B over 10 min, 70% to 100% B over 1 min, holdfor 4 min, 100% to 5% B over 1 s, hold for 5 min.

The CYN concentrations obtained (mg CYN L�1) were subse-quently standardised to each of the biomass parameters (chloro-phyll a concentration, dry weight, cell concentration, andbiovolume). For the batch experiments, the average values foreach experimental condition were calculated as the slope of alinear regression (p < 0.05) between CYN and each of the biomassparameters, including all samples from the 4-week experimentalperiod.

2.4. Statistical analysis

All statistical analyses were performed with SPSS Statisticssoftware (version 17.0.0, 2008).

3. Results

3.1. Effects of temperature and light-intensity on growth

The temperature-dependent growth experiments with A.

ovalisporum indicated that this species grows over a wide rangeof temperatures, from 15 8C (with a minimum growth rate of0.08 d�1) to 35 8C (with the maximum growth rate of 0.35 d�1)(Fig. 1). Extreme temperatures (10 8C and 40 8C) induced cell deathand net growth was not observed (Fig. 1). The growth rates of semi-continuous and batch experiments showed a similar trend,although this value was always slower for the batch experimentsthan for semi-continuous experiments; the batch growth rateswere an average of 21% less than the semi-continuous growth rates

Fig. 1. Temperature-dependent growth and Chl a content of A. ovalisporum UAM

290. Growth rates are represented by black circles and solid line in semi-continuous

experiments and by white circles and dashed line in batch experiments. Chl a

content is represented by white vertical bars in semi-continuous cultures and grey

vertical bars in batch cultures. Error bars indicate 95% confidence intervals.

Fig. 2. Light-dependent growth and Chl a content of A. ovalisporum UAM 289 grown

in batch. Growth rates are represented by black circles and solid line. Chl a is

represented by vertical bars. Error bars indicate 95% confidence intervals.

S. Cires et al. / Harmful Algae 10 (2011) 668–675 671

at equal temperatures. Chl a content closely followed the growth-rate curve within the 15–30 8C interval. In contrast, at 35 8C, inspite of the high growth rate recorded at this temperature, a strongdecrease in the content of Chl a (38% lower content than at 30 8C)was observed.

A. ovalisporum growing in batch cultures under differentirradiances (Fig. 2) showed growth rates ranging from 0.16 d�1

at the highest light-intensity assayed (340 mE photons m�2 s�1) to0.26 d�1 at 60 mE m�2 s�1. The lowest irradiance (15 mE m�2 s�1)resulted in lower growth rates but the maximum Chl a content. Atthe highest irradiance (340 mE m�2 s�1), the Chl a content was 30%lower than the maximum value.

3.2. Influence of strain and culture-medium on growth, CYN content

and release

The semi-continuous experiments carried out at a suboptimaltemperature of 30 8C and under 80 mE m�2 s�1 (Table S1)confirmed that, while strains UAM 289 and UAM 290 did notdiffer statistically (p > 0.05; t-test) in their growth rates or theirCYN content and release, the culture medium induced significantdifferences in the CYN content in UAM 290 (p < 0.05; t-test); theCYN content was 36% higher in BG110-grown culture than in Z8-grown culture. Similarly, the share of dissolved CYN (p < 0.05; t-test) was higher in BG110-grown culture (47.5% dissolved CYN)than in Z8-grown culture (31.7%).

3.3. Effects of temperature and light intensity on CYN content and

release

The effects of temperature on CYN content are shown in Fig. 3and Table 1. The biomass-standardised CYN content was obtained

Table 1Total CYN standardized to biomass and share of dissolved CYN of A. ovalisporum UAM

Temperature (8C) Growth regime Total CYN

(mg mg�1 DW) (mg mm�3)

15 Semi-continuous 2.24 � 0.29 0.79 � 0.10

Batch 1.77 � 0.18 0.62 � 0.06

20–30 rangea Semi-continuous 1.56 � 0.10 0.54 � 0.03

Batch 2.03 � 0.35 0.71 � 0.05

35 Semi-continuous 0.09 � 0.03 0.03 � 0.01

Values are expressed as mean � 95% confidence intervals for semi-continuous experiments

experiments.a Temperatures of 20, 25, 27.5 and 30 8C in semi-continuous experiments and 20, 27

in the batch experiments by linear regression (p < 0.05) betweentotal CYN and each of the biomass parameters. The statisticalanalysis revealed a clear influence of temperature on biomass-standardised CYN content for semi-continuous experiments(p < 0.05; one-way ANOVA) and the batch experiments(p < 0.05; one-way ANCOVA). The overall results from both typesof experiments showed that the most pronounced shift in CYNcontent took place at 35 8C, with a dramatic 25-fold decrease withrespect to the maximum concentration. Further, within the 15–30 8C interval, when the CYN content was standardised to biomass(DW, biovolume, cell number) (Table 1) there were only slightvariations in the CYN content of the semi-continuous experiments(1.9-fold) and of the batch experiments (1.3-fold). However, thesedifferences were increased when the CYN content was standar-dised to Chl a, with a maximum of 0.5 mg CYN mg�1 Chl a at 15 8Cthat was approximately 4-fold higher than the 20–30 8C average.

The maximum total-CYN content was reached at 15 8C in thesemi-continuous experiments (p < 0.05; Bonferroni post hoc test)and at 27.5 8C in the batch experiments (p < 0.05; t-test). Withinthis 15–30 8C interval, as detailed in Table 1, the maximum total-CYN content reached 2.2 mg mg�1 DW, which is equivalent to0.79 mg mm�3 and 87 fg cell�1.

Temperature induced significant differences on CYN release(Fig. 3), as demonstrated by statistical analysis both in semi-continuous experiments (p < 0.05; one-way ANOVA) and batchexperiments (p < 0.05; one-way ANCOVA). The post-hoc pair-wisecomparisons in the semi-continuous experiments (p < 0.05;Bonferroni test) revealed that extreme temperatures (15 8C and35 8C) showed a significantly higher share of dissolved CYN(accounting from 59% to 64% of the total CYN amount) thantemperatures between 20 and 30 8C, with 31% of dissolved CYN, asshown in Table 1. The results from the batch experiments

290 at different temperatures.

Dissolved CYN (% of total CYN)

(fg cell�1) (mg mg�1 Chl a)

86.64 � 11.05 0.51 � 0.10 63.50 � 3.81

68.47 � 6.83 0.40 � 0.05 60.20 � 5.10

44.69 � 4.75 0.12 � 0.02 31.16 � 3.31

78.87 � 5.62 0.16 � 0.01 22.55 � 1.23

2.31 � 0.65 0.01 � 0.01 58.72 � 13.04

and as estimated slope of regression lines (p < 0.05) �95% confidence intervals for batch

.5 and 30 8C in batch experiments.

Fig. 3. Total CYN content and share of dissolved CYN of A. ovalisporum UAM 290 at

different temperatures. Total CYN per unit of biovolume is represented by white

vertical bars in semi-continuous experiments and by grey vertical bars in batch

experiments. Dissolved CYN is represented by black triangles and solid line in semi-

continuous experiments and by white triangles and dashed line in batch-

experiments. Error bars indicate 95% confidence intervals.

Fig. 4. Total CYN content and share of dissolved CYN of A. ovalisporum UAM 289

grown in batch under different light intensities. Total CYN per unit of biovolume is

represented by vertical bars. Dissolved CYN is represented by black triangles and

solid line. Error bars indicate 95% confidence intervals.

Fig. 5. Relationship between total CYN content and growth rates (A) and between

share of dissolved CYN and growth rates (B) in A. ovalisporum under all

temperatures and light intensities assayed. In both graphs, results from

temperature experiments are represented by black circles and results from

light-intensity experiments are represented by white triangles. A regression line

(y = 1.90x � 2.60; R2 = 0.93; n = 4) is plotted in (A) for total CYN contents vs growth

rates in light-intensity experiments.

S. Cires et al. / Harmful Algae 10 (2011) 668–675672

supported this finding, but with a slightly lower amount ofdissolved CYN (approximately 22%) for the 20–30 8C interval.

Changes in the light-intensity influenced the CYN content in A.

ovalisporum (Fig. 4), as revealed by the statistical analysis showingsignificant differences between all the intensities assayed(p < 0.05; one-way ANCOVA). Nevertheless, the CYN contentwas within the same order of magnitude over the whole light-intensity range assayed. At a range of 15–60 mE m�2 s�1, the total-CYN content per biovolume increased 2-fold, showing themaximum total-CYN recorded with this strain of 6.4 mg mg�1

Table 2Total CYN standardized to biomass and share of dissolved CYN of A. ovalisporum UAM

Light intensity (mE m�2 s�1) Total CYN

(mg mg�1 DW) (mg mm�3) (

15 3.49 � 0.73 1.25 � 0.14

60 6.37 � 0.56 2.43 � 0.35 1

200 2.38 � 0.60 0.90 � 0.16

340 1.32 � 0.15 0.58 � 0.07

Values are expressed as estimated slope of regression lines (p < 0.05) �95% confidence

differences were found (p > 0.05; one-way ANCOVA).

DW (Table 2). In contrast, higher light intensities of 200 and340 mE m�2 s�1, considered growth-saturating intensities (seeFig. 2), triggered a remarkable decrease of CYN content, whichwas 4-fold lower than the maximum value.

The share of dissolved CYN was not significantly affected bylight-intensity (Fig. 4), since no statistical differences were found(p > 0.05; one-way ANCOVA) between slopes of dissolved-CYN vs

total-CYN regression lines at different light intensities. This fact ledus to group all dissolved CYN values from light-intensityexperiments in a global regression, resulting in an average shareof 25% of total CYN in the dissolved fraction (Table 2).

3.4. CYN content and release related to cyanobacterial growth

Total-CYN content showed no global relationship with growthrates, as shown by Fig. 5a, even though a positive linear

289 grown in batch under different light intensities.

Dissolved CYN (% of total CYN)

fg cell�1) (mg mg�Chl a)

97.65 � 10.67 0.28 � 0.05 25.17 � 3.71

90.62 � 26.61 0.53 � 0.04

70.75 � 12.15 0.34 � 0.06

45.20 � 5.75 0.16 � 0.02

intervals. Dissolved CYN is averaged for all the light intensities since no statistical

S. Cires et al. / Harmful Algae 10 (2011) 668–675 673

relationship can be established between these two parameterswhen looking at the light-intensity experiments alone, as growthrates increased in parallel with CYN content.

The share of dissolved CYN was not linearly correlated togrowth rate, although a trend can be observed in the scatter plot(Fig. 5b), with the highest proportion of dissolved CYN located atboth extremes of the growth rates range (below 0.1 d�1 and above0.3 d�1). Most data within moderate growth rates from 0.1 to0.3 d�1 showed an average dissolved-CYN share of 20–35% of totalCYN, independent of the environmental condition affecting growthvariations.

4. Discussion

To date, A. ovalisporum isolates have been reported fromwaterbodies in Australia, Florida, Israel and Spain (Stuken et al.,2009, and references within). Laboratory experiments on theecophysiology of this species have been conducted with strainsfrom Israel (Hadas et al., 2002) and, more recently, from Spain(Mehnert et al., 2010; and the present study). As a whole, thesereports demonstrate maximum growth rates of 0.2–0.36 d�1 for A.

ovalisporum, which agrees in situ estimations in Lake Kinneret(Pollingher et al., 1998), and in the range of other CYN producerslike A. flos-aquae (Preussel et al., 2009) or C. raciborskii in the field(Padisak and Istvanovics, 1997). In our study, A. ovalisporum grewat a temperature range of 15–35 8C; a range similar to thoseoccurring in thermally-stratified Mediterranean waterbodieswhere this species is naturally present (Pollingher et al., 1998;Gkelis et al., 2005). Temperature optima differed slightly betweenthe ecotypes from Israel (26–30 8C) (Hadas et al., 2002) and Spain(32.8 8C) (Mehnert et al., 2010), similar to the 28–31 8C found forisolates of the invasive C. raciborskii from various latitudes (Briandet al., 2004). We observed the maximum growth of A. ovalisporum

at a moderate light of 60 mE m�2 s�1; this is not surprising becauseHadas et al. (2002) measured the maximum photosyntheticefficiency of Israeli strains at 40–100 mE m�2 s�1, resembling lightincident at 1–5 m depths during the 1994 bloom in Lake Kinneret.Furthermore, these authors found light saturation points (Ik) of 40–100 mE m�2 s�1 for A. ovalisporum; these saturation points aremidway between those estimated for C. raciborskii (Ik = 15–26)(Briand et al., 2004) and Microcystis aeruginosa

(Ik = 155 mE m�2 s�1) (Coles and Jones, 2000) or Anabaena flos-

aquae (Ik = 200 mE m�2 s�1) (De Nobel et al., 1998).The demonstrated tolerance to a wide range of temperatures

and light intensities, together with its buoyancy regulation ability(Porat et al., 2001) might facilitate A. ovalisporum survival indiverse habitats. This fact contrasts with the low number (althoughincreasing in the last years) of blooms reported for this species(Banker et al., 1997; Shaw et al., 1999; Quesada et al., 2006;Messineo et al., 2010). In addition to warm water temperatures(Pollingher et al., 1998), other specific conditions might benecessary to trigger those infrequent blooms, such as high pHand alkalinity, which enhances the experimentally proven use ofHCO3

� by A. ovalisporum (Hadas et al., 2002), or low N:P ratios(Pollingher et al., 1998; Quesada et al., 2006) which favournitrogen-fixers.

We confirmed a high total-CYN content in A. ovalisporum fromSpain of up to 6.4 mg mg�1 DW, 2.4 mg mm�3, 190 fg cell�1 or0.5 mg mg�1 Chl a, similar to the estimated CYN/biomass ratios inthe Arcos reservoir (Spain) (Quesada et al., 2006), in the range ofCYN contents in other A. ovalisporum strains, A. flos-aquae or C.

raciborskii, and clearly above the other CYN-producers (Table S2;Hawkins et al., 1997; Li et al., 2001b; McGregor et al., 2011;Preussel et al., 2006; Seifert et al., 2007; Spoof et al., 2006). Theconstitutive CYN production by all reported strains of A.

ovalisporum from different latitudes contrasts with the geographi-

cal separation between CYN-producing and non-producing strainsof C. raciborskii (Haande et al., 2008), the restricted location of someCYN-producers, such as Anabaena lapponica or Umezakia natans, orthe low proportion of toxin-producing strains of the cosmopolitancyanobacteria A. flos-aquae, Raphidiopsis curvata and Raphidiopsis

mediterranea, thus supporting the use of A. ovalisporum as a modelspecies for studies on CYN production and release.

A. ovalisporum produced CYN from 15 8C to 35 8C, withtemperature inducing only 2-fold variations within the 15–30 8Cinterval, and a dramatic decrease up to an almost total inhibition at35 8C. In spite of a low growth rate, a high CYN content wasobserved at low temperatures, as already observed in A. flos-aquae

(Preussel et al., 2009). Similar to our results, CYN production wascompletely inhibited at 35 8C in C. raciborskii (Saker and Griffiths,2000), a behaviour that may be related to the loss of activity over35 8C in the amidinotransferase CyrA that is involved in CYNsynthesis in C. raciborskii (Muenchhoff et al., 2010). Hypothetically,the amidinotransferase A (encoded by gene aoaA) involved in CYNsynthesis by A. ovalisporum (Shalev-Alon et al., 2002) might followa similar trend, although no data is available on temperature-related stability of this enzyme to date.

The light-dependent CYN content in A. ovalisporum wascharacterised by 4-fold variations within a light-irradiance range(15–200 mE m�2 s�1) similar to that measured at 0.5–9.5 m depthsduring the 1994 bloom in Lake Kinneret (Hadas et al., 2002).Considering that aoaA and aoaC transcription levels are onlyreversibly affected by light irradiance (Shalev-Malul et al., 2008), apost-transcriptional regulation is more likely to occur in A.

ovalisporum. The light irradiance induced different responses inother species, not having a significant influence in A. flos-aquae

(Preussel et al., 2009) whereas being negatively correlated to CYNproduction in C. raciborskii (Dyble et al., 2006).

Besides light and temperature, nutrients like sulphate andphosphorus may also influence CYN production in A. ovalisporum

(Bacsi et al., 2006; Bar-Yosef et al., 2010) thereby explaining thedifferent CYN contents we found in the same strain under twodifferent culture media (BG110 and Z8) with different nutrientconcentrations (Rippka et al., 1979; Kotai, 1972). In the particularcase of phosphorous, the results published to date are contradic-tory. Bacsi et al. (2006) reported a CYN decrease under phosphoruslimitation, whereas Bar-Yosef et al. (2010) established a CYNincrease in A. ovalisporum grown under phosphorus-limitingconditions. In our case, both CYN content and release were higherunder the medium with lower phosphorus concentration (BG110),somehow resembling findings from Bar-Yosef and co-workers.However, the increased differences between the culture media weused (BG110 and Z8) in consideration of additional macro andmicro-nutrients other than phosphorous lead us to take thesefindings with caution.

Information about extracellular CYN release in differentcyanobacterial species is still scarce. All the previous experimentsdemonstrated a conspicuous release, of up to 40–58% of dissolvedCYN during exponential growth in semi-continuous cultures of A.

flos-aquae (Preussel et al., 2009) and above 50% during thestationary phase of batch-grown C. raciborskii (Hawkins et al.,2001; Saker and Griffiths, 2000), although the results in the latterare influenced by the use of batch cultures in which the dissolvedCYN might have accumulated due to a lack of degradation. Wefound an even higher CYN release in A. ovalisporum grown under asemi-continuous regime, with a maximum of 58–63% of dissolvedCYN under extreme temperatures (35 8C and 15 8C), almost 2 timeshigher than the 20–30 8C average, these shifts being mainly drivenby temperature without an influence of light irradiance. Incontrast, CYN release in A. flos-aquae was not significantlyinfluenced either by temperature or by light intensity (Preusselet al., 2009), notably the temperature and light ranges assayed in

S. Cires et al. / Harmful Algae 10 (2011) 668–675674

that study (16–25 8C; 10–60 mE m�2 s�1) were considerablynarrower than those of our experimental setup.

To date, the active or passive nature of CYN transport acrosscellular membranes remains unsolved. We estimated an intracel-lular single-cell concentration of CYN to be 1000 times higher thanthe extracellular one (estimated extracellular concentrationranging from 0.1% to 1% of intracellular concentration in thedifferent experiments, data not shown), suggesting that release isfavored by gradient and thus, is not necessarily an energy-consuming process. Therefore, both passive diffusion and facilitateddiffusion are hypothetical candidates for CYN transport, althoughno evidence supporting these processes has been reported inliterature to date. Furthermore, as cell membrane stability was notmonitored either in any of the previously reported experiments onCYN-release or in the present study, differences in permeability ofthe membrane influencing the transport cannot be discarded.Moreover, Mihali et al. (2008) previously postulated the existenceof the sodium ion-driven CYN transporter CyrK in C. raciborskii,encoded by the gene cyrK belonging to the CYN-synthesis cluster.The product encoded by cyrK was similar to a family of toxiccompound extrusion proteins (NorM) within the MATE-typeproteins, transporters of organic cationic drugs in bacteria andsecondary metabolites and xenobiotics in plants (Omote et al.,2006), substances somehow related to CYN as it is a cationicsecondary metabolite (Ohtani et al., 1992). CyrK orthologous havebeen also found in A. flos-aquae (Stuken and Jakobsen, 2010) andOscillatoria sp. PCC6506 (Mazmouz et al., 2010), even though itsexistence in A. ovalisporum still remains unknown.

Whereas MCYST production is considered to be positivelycorrelated with growth regardless of the stressor acting (Orr andJones, 1998), an equivalent trend for CYN was not found for anyCYN-producer up to date (Saker and Neilan, 2001; Dyble et al.,2006; Preussel et al., 2009; and the present study). In the case ofthe light-intensity experiments in our study, a linear relationship(n = 4) between CYN contents and growth rates can be observed(see Fig. 5a), although we interpreted this trend as an indirectrelationship because of the effect of light intensity on growth rates,as previously described (see Fig. 2).

Extracellular CYN release was negatively correlated togrowth in the 0.11–0.20 d�1 growth range in A. flos-aquae

(Preussel et al., 2009). This result contrasts with the global trendwe observed in A. ovalisporum for a wider growth range of 0.08–0.35 d�1. In fact, we observed a slightly variable CYN release of20–35% within 0.1–0.3 d�1, growth rates in the range of in situ

estimations for field populations (Pollingher et al., 1998), whichincreased below 0.1 d�1 and above 0.3 d�1. Further, Preusselet al. (2009) also suggested a stress-related increase in CYNrelease, a hypothesis we cannot discard completely because inour study, points with more than 50% of dissolved CYN (seeFig. 5b) occur at 15 8C, where growth was clearly reduced andthus cultures may be stressed, and at 35 8C, a condition that, inspite of the high growth rate recorded, may be close to a stressthreshold, as suggested by a decrease in Chl a and CYN contentindicating an altered metabolism.

In view of our results, CYN is highly likely to occur when A.

ovalisporum is present in a given waterbody and, in that case,only 5000 cells mL�1, 0.5 mm3 L�1 or 2 mg Chl a L�1 of thisspecies, still far from bloom formation, may be sufficient toreach the recommended drinking water guideline of1 mg CYN L�1 (Humpage and Falconer, 2003). Therefore, bio-mass-based guideline values specifically developed for A.

ovalisporum are needed, similar to those proposed by QueenslandHealth Services (Australia) (Quesada et al., 2006) but (to ourknowledge) not present in other legislations yet. Furthermore,future normative should pay special attention to recreationaluses of water as CYN in the dissolved fraction and thus in direct

contact with users may represent a 20–35% of the totalconcentration even before bloom decay.

5. Conclusions

As a whole, we demonstrate A. ovalisporum to be an ecologicallyplastic species producing CYN in the range of mg mg�1 DW overbroad temperature and light ranges that naturally occur through-out the annual cycle of the current and potential habitats of thisspecies. In addition, we estimated that 20–35% of extracellular CYNis to be expected within typical growth ranges of this organism inthe field, suddenly increasing to more than 50% under extremetemperatures and/or growth rates. Because of the genetichomogeneity demonstrated for this species worldwide (Stukenet al., 2009), which is also supported by the high similarities wefound between UAM 289 and UAM 290, both genetically (100%similarity on cpcBA-IGS and 16S rRNA gene) and in terms of CYNproduction and release, our findings may be easily applicable toother A. ovalisporum strains and field populations.

Acknowledgements

Samuel Cires and Lars Wormer were supported by a FPU grantfrom Ministerio de Ciencia e Innovacion (MICINN, Spain). Thisstudy was partially funded by the German Ministry of Educationand Research and Kompetenzzetrum Wasser Berlin with financialsupport from Veolia Water and Berliner Wasserbetriebe. Ouracknowledgment to all members of Molecular Ecology Group fromIGB (Stechlin, Germany), especially to Grit Mehnert, FranziskaLeunert and Maren Lentz for their valuable help with semi-continuous cultures, and to Dr. Andreas Ballot for his teaching ofmolecular techniques. We would also like to thank two anonymousreviewers for their helpful comments.[SS]

Appendix A. Supplementary data

Supplementary data associated with this article can be found, in

the online version, at doi:10.1016/j.hal.2011.05.002.

References

Bacsi, I., Vasas, G., Suranyi, G., Hamvas, M., Mathe, C., Toth, E., Grigorszy, I., Gaspar,A., Toth, S., Borbely, G., 2006. Alteration of cylindrospermopsin production insulfate- or phosphate-starved cyanobacterium Aphanizomenon ovalisporum.FEMS Microbiol. Lett. 259 (2), 303–310.

Ballot, A., Dadheech, P.K., Haande, S., Krienitz, L., 2008. Morphological and phylo-genetic analysis of Anabaenopsis abijatae and Anabaenopsis elenkinii (Nostocales,Cyanobacteria) from tropical inland water bodies. Microb. Ecol. 55 (4), 608–618.

Ballot, A., Fastner, J., Wiedner, C., 2010. Paralytic shellfish poisoning toxin-produc-ing Cyanobacterium Aphanizomenon gracile in Northeast Germany. Appl. Envi-ron. Microbiol. 76 (4), 1173–1180.

Banker, R., Carmeli, S., Hadas, O., Teltsch, B., Porat, R., Sukenik, A., 1997. Identifica-tion of cylindrospermopsin in Aphanizomenon ovalisporum (Cyanophyceae)isolated from Lake Kinneret. Isr. J. Phycol. 33 (4), 613–616.

Bar-Yosef, Y., Sukenik, A., Hadas, O., Viner-Mozzini, Y., Kaplan, A., 2010. Enslave-ment in the water body by toxic Aphanizomenon ovalisporum, inducing alkalinephosphatase in phytoplanktons. Curr. Biol. 20, 1–5.

Blahova, L., Oravec, M., Marsalek, B., Sejnohova, L., Simek, Z., Blaha, L., 2009. The firstoccurrence of the cyanobacterial alkaloid toxin cylindrospermopsin in theCzech Republic as determined by immunochemical and LC/MS methods. Tox-icon 53 (5), 519–524.

Briand, J.F., Leboulanger, C., Humbert, J.F., Bernard, C., Dufour, P., 2004. Cylindros-permopsis raciborskii (Cyanobacteria) invasion at mid-latitudes: selection, widephysiological tolerance, or global warming? J. Phycol. 40 (2), 231–238.

Brient, L., Lengronne, M., Bormans, M., Fastner, J., 2009. First occurrence of cylin-drospermopsin in freshwater in France. Environ. Toxicol. 24 (4), 415–420.

Burns, J., Williams, C., Champan, A., 2002. Cyanobacteria and their toxins in Floridasurface waters. In: Johnson, D., Harbison, R.D. (Eds.), Abstracts of the MeetingHealth Effects of Exposure to Cyanobacteria Toxins: State of the Science,Saratoga, FL, pp. 16–21.

S. Cires et al. / Harmful Algae 10 (2011) 668–675 675

Codd, G.A., Morrison, L.F., Metcalf, J.S., 2005. Cyanobacterial toxins: risk manage-ment for health protection. Toxicol. Appl. Pharmacol. 203 (3), 264–272.

Coles, J.F., Jones, R.C., 2000. Effect of temperature on photosynthesis-light responseand growth of four phytoplankton species isolated from a tidal freshwater river.J. Phycol. 36 (1), 7–16.

De Nobel, W.T., Matthijs, H.C.P., Von Elert, E., Mur, L.R., 1998. Comparison of thelight-limited growth of the nitrogen-fixing cyanobacteria Anabaena and Apha-nizomenon. New Phytol. 138 (4), 579–587.

Dyble, J., Tester, P.A., Litaker, R.W., 2006. Effects of light intensity on cylindros-permopsin production in the cyanobacterial HAB species Cylindrospermopsisraciborskii. Afr. J. Mar. Sci. 28 (2), 309–312.

Gkelis, S., Moustaka-Gouni, M., Sivonen, K., Lanaras, T., 2005. First report of thecyanobacterium Aphanizomenon ovalisporum Forti in two Greek lakes andcyanotoxin occurrence. J. Plankton Res. 27 (12), 1295–1300.

Griffiths, D.J., Saker, M.L., 2003. The Palm Island mystery disease 20 years on: areview of research on the cyanotoxin cylindrospermopsin. Environ. Toxicol. 18(2), 78–93.

Haande, S., Rohrlack, T., Ballot, A., Roberg, K., Skulberg, R., Beck, M., Wiedner, C.,2008. Genetic characterisation of Cylindrospermopsis raciborskii (Nostocales,Cyanobacteria) isolates from Africa and Europe. Harmful Algae 7 (5), 692–701.

Hadas, O., Pinkas, R., Malinsky-Rushansky, N., Shalev-Alon, G., Delphine, E., Berner,T., Sukenik, A., Kaplan, A., 2002. Physiological variables determined underlaboratory conditions may explain the bloom of Aphanizomenon ovalisporumin Lake Kinneret. Eur. J. Phycol. 37 (2), 259–267.

Hawkins, P.R., Chandrasena, N.R., Jones, G.J., Humpage, A.R., Falconer, I.R., 1997.Isolation and toxicity of Cylindrospermopsis raciborskii from an ornamental lake.Toxicon 35 (3), 341–346.

Hawkins, P.R., Putt, E., Falconer, I., Humpage, A., 2001. Phenotypical variation in atoxic strain of the phytoplankter Cylindrospermopsis raciborskii (Nostocales,Cyanophyceae) during batch culture. Environ. Toxicol. 16 (6), 460–467.

Humpage, A.R., Falconer, I.R., 2003. Oral toxicity of the cyanobacterial toxin cylin-drospermopsin in male Swiss albino mice: determination of no observedadverse effect level for deriving a drinking water guideline value. Environ.Toxicol. 18 (2), 94–103.

Kinnear, S.H.W., Duivenvoorden, L.J., Fabbro, L.D., 2007a. Sublethal responses inMelanoides tuberculata following exposure to Cylindrospermopsis raciborskiicontaining cylindrospermopsin. Harmful Algae 6 (5), 642–650.

Kinnear, S.H.W., Fabbro, L.D., Duivenvoorden, L.J., Hibberd, E.M.A., 2007b. Multiple-organ toxicity resulting from cylindrospermopsin exposure in tadpoles of thecane toad (Bufo marinus). Environ. Toxicol. 22 (6), 550–558.

Kokocinski, M., Dziga, D., Spoof, L., Stefaniak, K., Jurczak, T., Mankiewicz-Boczek, J.,Meriluoto, J., 2009. First report of the cyanobacterial toxin cylindrospermopsin inthe shallow, eutrophic lakes of western Poland. Chemosphere 74 (5), 669–675.

Kotai, J., 1972. Instructions For Preparation of Modified Nutrient Solution Z8 forAlgae. Norwegian Institute for Water Research, Oslo.

Li, R., Carmichael, W.W., Brittain, S., Eaglesham, G.K., Shaw, G.R., Mahakhant, A.,Noparatnaraporn, N., Yongmanitchai, W., Kaya, K., Watanabe, M.M., 2001a.Isolation and identification of the cyanotoxin cylindrospermopsin and deoxy-cylindrospermopsin from a Thailand strain of Cylindrospermopsis raciborskii(Cyanobacteria). Toxicon 39 (7), 973–980.

Li, R.H., Carmichael, W.W., Brittain, S., Eaglesham, G.K., Shaw, G.R., Liu, Y.D.,Watanabe, M.M., 2001b. First report of the cyanotoxins cylindrospermopsinand deoxy-cylindrospermopsin from Raphidiopsis curvata (Cyantobacteria). J.Phycol. 37 (6), 1121–1126.

Marker, A.F., Nush, E.A., Rai, H.B.R., 1980. The measurement of photosyntheticpigments in freshwaters and standardization of methods: conclusions andrecommendations. Arch. Hydrobiol. 14, 91–106.

Mazmouz, R., Chapuis-Hugon, F., Mann, S., Pichon, V., Mejean, A., Ploux, O., 2010.Biosynthesis of cylindrospermopsin and 7-epicylindrospermopsin in Oscilla-toria sp. strain PCC 6506: identification of the cyr gene cluster and toxinanalysis. Appl. Environ. Microbiol. 76 (15), 4943–4949.

McGregor, G.B., Sendall, B.C., Hunt, L.T., Eaglesham, G.K., 2011. Report of thecyanotoxins cylindrospermopsin and deoxy-cylindrospermopsin from Raphi-diopsis mediterranea Skuja (Cyanobacteria/Nostocales). Harmful Algae 10, 402–410.

Mehnert, G., Leunert, F., Cires, S., Johnk, K.D., Rucker, J., Nixdorf, B., Wiedner, C.,2010. Competitiveness of invasive and native cyanobacteria from temperatefreshwaters under various light and temperature conditions. J. Plankton Res. 32(7), 1009–1021.

Messineo, V., Melchiorre, S., Di Corcia, A., Gallo, P., Bruno, M., 2010. SeasonalSuccession of Cylindrospermopsis raciborskii and Aphanizomenon ovalisporumBlooms with Cylindrospermopsin Occurrence in the Volcanic Lake Albano,Central Italy. Environ. Toxicol. 25 (1), 18–27.

Mihali, T.K., Kellmann, R., Muenchhoff, J., Barrow, K.D., Neilan, B.A., 2008. Charac-terization of the gene cluster responsible for cylindrospermopsin biosynthesis.Appl. Environ. Microbiol. 74 (3), 716–722.

Muenchhoff, J., Siddiqui, K.S., Poljak, A., Raftery, M.J., Barrow, K.D., Neilan, B.A., 2010.A novel prokaryotic L-arginine:glycine amidinotransferase is involved in cylin-drospermopsin biosynthesis. FEBS J. 277 (18), 3844–3860.

Mur, L.R., Skulberg, O.M., Utkilen, H., 1999. Cyanobacteria in the environment. In:Chorus, I., Bartram, J. (Eds.), Toxic Cyanobacteria in Water: A Guide of TheirPublic Health Consequences, Monitoring and Management. E & FN Spoon,London and New York, pp. 15–40.

Nogueira, I.C.G., Saker, M.L., Pflugmacher, S., Wiegand, C., Vasconcelos, V.M., 2004.Toxicity of the cyanobacterium Cylindrospermopsis raciborskii to Daphniamagna. Environ. Toxicol. 19 (5), 453–459.

Ohtani, I., Moore, R.E., Runnegar, M.T.C., 1992. Cylindrospermopsin: a potenthepatotoxin from the blue-green alga Cylindrospermopsis raciborskii. J. Am.Chem. Soc. 114 (20), 7941–7942.

Omote, H., Hiasa, M., Matsumoto, T., Otsuka, M., Moriyama, Y., 2006. The MATEproteins as fundamental transporters of metabolic and xenobiotic organiccations. Trends Pharmacol. Sci. 27 (11), 587–593.

Orr, P.T., Jones, G.J., 1998. Relationship between microcystin production and celldivision rates in nitrogen-limited Microcystis aeruginosa cultures. Limnol. Ocea-nogr. 43 (7), 1604–1614.

Padisak, J., Istvanovics, V., 1997. Differential response of blue-green algal groups tophosphorus load reduction in a large shallow lake: Balaton, Hungary. In:Williams, W.D., Sladeckova, A. (Eds.), International Association of Theoreticaland Applied Limnology, Vol. 26, Pt 2, pp. 574–580.

Pearson, L., Mihali, T., Moffitt, M., Kellmann, R., Neilan, B., 2010. On the chemistry,toxicology and genetics of the cyanobacterial toxins, microcystin, nodularin,saxitoxin and cylindrospermopsin. Mar. Drugs 8 (5), 1650–1680.

Pollingher, U., Hadas, O., Yacobi, Y.Z., Zohary, T., Berman, T., 1998. Aphanizomenonovalisporum (Forti) in Lake Kinneret, Israel. J. Plankton Res. 20 (7), 1321–1339.

Porat, R., Teltsch, B., Perelman, A., Dubinsky, Z., 2001. Diel buoyancy changes by thecyanobacterium Aphanizomenon ovalisporum from a shallow reservoir. J. Plank-ton Res. 23 (7), 753–763.

Preussel, K., Stuken, A., Wiedner, C., Chorus, I., Fastner, J., 2006. First report oncylindrospermopsin producing Aphanizomenon flos-aquae (Cyanobacteria) iso-lated from two German lakes. Toxicon 47 (2), 156–162.

Preussel, K., Wessel, G., Fastner, J., Chorus, I., 2009. Response of cylindrospermopsinproduction and release in Aphanizomenon flos-aquae (Cyanobacteria) to varyinglight and temperature conditions. Harmful Algae 8 (5), 645–650.

Quesada, A., Moreno, E., Carrasco, D., Paniagua, T., Wormer, L., De Hoyos, C., Sukenik,A., 2006. Toxicity of Aphanizomenon ovalisporum (Cyanobacteria) in a Spanishwater reservoir. Eur. J. Phycol. 41 (1), 39–45.

Rippka, R., Deruelles, J., Waterbury, J.B., Herdman, M., Stanier, R.Y., 1979. Genericassignments, strain histories and properties of pure cultures of cyanobacteria. J.Gen. Microbiol. 111 (March), 1–61.

Saker, M.L., Griffiths, D.J., 2000. The effect of temperature on growth and cylin-drospermopsin content of seven isolates of Cylindrospermopsis raciborskii (Nos-tocales, Cyanophyceae) from water bodies in northern Australia. Phycologia 39(4), 349–354.

Saker, M.L., Neilan, B.A., 2001. Varied diazotrophies, morphologies, and toxicities ofgenetically similar isolates of Cylindrospermopsis raciborskii (Nostocales, Cyano-phyceae) from northern Australia. Appl. Environ. Microbiol. 67 (4), 1839–1845.

Seifert, M., McGregor, G., Eaglesham, G., Wickramasinghe, W., Shaw, G., 2007. Firstevidence for the production of cylindrospermopsin and deoxy-cylindrosper-mopsin by the freshwater benthic cyanobacterium Lyngbya wollei (Farlow exGornont) Speziale and Dyck. Harmful Algae 6 (1), 73–80.

Shalev-Alon, G., Sukenik, A., Livnah, O., Schwarz, R., Kaplan, A., 2002. A novel geneencoding amidinotransferase in the cylindrospermopsin producing cyanobac-terium Aphanizomenon ovalisporum. FEMS Microbiol. Lett. 209 (1), 87–91.

Shalev-Malul, G., Lieman-Hurwitz, J., Viner-Mozzini, Y., Sukenik, A., Gaathon, A.,Lebendiker, M., Kaplan, A., 2008. An AbrB-like protein might be involved in theregulation of cylindrospermopsin production by Aphanizomenon ovalisporum.Environ. Microbiol. 10 (4), 988–999.

Shaw, G.R., Seawright, A.A., Moore, M.R., Lam, P.K.S., 2000. Cylindrospermopsin, acyanobacterial alkaloid: evaluation of its toxicologic activity. Ther. Drug Monit.22 (1), 89–92.

Shaw, G.R., Sukenik, A., Livne, A., Chiswell, R.K., Smith, M.J., Seawright, A.A., Norris,R.L., Eaglesham, G.K., Moore, M.R., 1999. Blooms of the cylindrospermopsincontaining cyanobacterium Aphanizomenon ovalisporum (Forti) in newly con-structed lakes, Queensland, Australia. Environ. Toxicol. 14 (1), 167–177.

Spoof, L., Berg, K.A., Rapala, J., Lahti, K., Lepisto, L., Metcalf, J.S., Codd, G.A., Meriluoto,J., 2006. First observation of cylindrospermopsin in Anabaena lapponica isolatedfrom the boreal environment (Finland). Environ. Toxicol. 21 (6), 552–560.

Stuken, A., Campbell, R.J., Quesada, A., Sukenik, A., Dadheech, P.K., Wiedner, C., 2009.Genetic and morphologic characterization of four putative cylindrospermopsinproducing species of the cyanobacterial genera Anabaena and Aphanizomenon. J.Plankton Res. 31 (5), 465–480.

Stuken, A., Jakobsen, K.S., 2010. The cylindrospermopsin gene cluster of Aphani-zomenon sp strain 10E6: organization and recombination. Microbiology 156,2438–2451.

Torokne, A., Asztalos, M., Bankine, M., Bickel, H., Borbely, G., Carmeli, S., Codd, G.A.,Fastner, J., Huang, Q., Humpage, A., Metcalf, J.S., Rabai, E., Sukenik, A., Suranyi, G.,Vasas, G., Weiszfeiler, V., 2004. Interlaboratory comparison trial on cylindros-permopsin measurement. Anal. Biochem. 332 (2), 280–284.

Utermohl, H., 1958. Zur Vervollkommung der quantitativen Phytoplankton-Meth-odik Mitt. Int. Ver. Limnol. 9, 1–38.

Wormer, L., Cires, S., Carrasco, D., Quesada, A., 2008. Cylindrospermopsin is notdegraded by co-occurring natural bacterial communities during a 40-day study.Harmful Algae 7 (2), 206–213.

Wormer, L., Carrasco, D., Cires, S., Quesada, A., 2009. Advances in solid phaseextraction of the cyanobacterial toxin cylindrospermopsin. Limnol. Oceanogr.7, 568–575.

Wormer, L., Huerta-Fontela, M., Cires, S., Carrasco, D., Quesada, A., 2010. Naturalphotodegradation of the cyanobacterial toxins microcystin and cylindrosper-mopsin. Environ. Sci. Technol. 44 (8), 3002–3007.

Yilmaz, M., Phlips, E.J., Szabo, N.J., Badylak, S., 2008. A comparative study of Floridastrains of Cylindrospermopsis and Aphanizomenon for cylindrospermopsin pro-duction. Toxicon 51 (1), 130–139.