Selective method for cyanobacterial bloom removal:hydraulic jet cavitation experience

Daniel Jancula • Premysl Mikula • Blahoslav Marsalek •

Pavel Rudolf • Frantisek Pochyly

Received: 13 August 2012 / Accepted: 13 June 2013 / Published online: 22 June 2013� Springer Science+Business Media Dordrecht 2013

Abstract The aim of this study was to investigate the suitability of hydraulic jet cavi-

tation as a method for cyanobacterial water-bloom management. Effects of cavitation were

studied on laboratory culture of the cyanobacterium Microcystis aeruginosa, on a culture

of a green alga Chlorella kessleri (as a non-target species) as well as on a real cyano-

bacterial biomass with Microcystis sp. as a dominant species. Our results suggested that the

cavitation treatment of cyanobacteria is capable of causing the disintegration of their gas

vesicles. Using this treatment, up to 99 % removal efficiency of cyanobacteria was

achieved. Moreover, no effect on cyanobacterial membrane integrity or metabolic activity

was detected by flow cytometry; thus, hydraulic cavitation seems to be harmless from the

viewpoint of possible release of cyanotoxins into the water column. The green algae (here

C. kessleri) were not affected negatively by the cavitation, and thus, they may still act as

the natural nutrient competitors of cyanobacteria in lakes, ponds or reservoirs treated by

cavitation.

Keywords Esterase activity � Membrane integrity � Bioassay � Ultrastructure �Flow cytometry

Introduction

Cyanobacterial (blue-green algal) proliferation is among the most threatening conse-

quences of freshwater pollution caused by humans. The development of cyanobacterial

water blooms occurs in reservoirs, lakes and ponds across the world (Lehman et al. 2005;

D. Jancula (&) � P. Mikula � B. MarsalekInstitute of Botany, Academy of Sciences of the Czech Republic, Lidicka 25/27, 60200 Brno,Czech Republice-mail: jancula@sinice.cz

P. Rudolf � F. PochylyV. Kaplan Department of Fluid Engineering, Faculty of Mechanical Engineering, Brno Universityof Technology, Technicka 2896/2, 61669 Brno, Czech Republic

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Aquacult Int (2014) 22:509–521DOI 10.1007/s10499-013-9660-7

Kabzinski et al. 2000; Kemp and John 2006). Cyanobacteria possess several competitive

advantages over other phytoplankton taxa. Most cyanobacteria can regulate their vertical

position in a water column thanks to gas vacuoles (Hayes and Walsby 1986), and they can

absorb and utilize bicarbonate (Koropatkin et al. 2007) and convert gaseous nitrogen into

ammonia via nitrogen fixation (Haselkorn 1986). Cyanobacteria are also well-known

producers of toxins which can be extremely toxic for a broad spectrum of organisms

including humans (Costa et al. 2009; Smith et al. 2008). Cyanotoxins may not only neg-

atively affect natural aquatic ecosystems but also the health of human beings utilizing

water from lakes, reservoirs and dams for everyday requirements (Douma et al. 2010). To

guarantee drinking water safety, a chronic tolerable daily intake (TDI) of 0.04 lg kg-1

body weight has been derived for microcystin-LR (Carmichael et al. 2001), and the World

Health Organization (WHO) also recommends a provisional guideline value for this

compound in drinking water being 1 lg L-1 (WHO 1998).

Various methods and techniques have been developed in order to decrease the abun-

dance of nuisance phytoplankton species in the water bodies (Gregor et al. 2008; Marsalek

et al. 2012). Although nutrient removal can warrant positive long-term effects leading to

the reduction in the trophic state and thus to the reduction in nuisance phytoplankton

species (namely cyanobacteria), significant restriction of nutrient inputs to the surface

waters is almost impossible and unavailable for most areas across the world due to con-

temporary economical limitations nowadays (Jancula and Marsalek 2011).

One of the most common and the cheapest methods of harmful phytoplankton bloom

control is the application of chemicals. Although their effects on cyanobacteria are usually

fast and effective (at least for a short period), their toxicity to non-target species and per-

sistence in the environment represent limitations of their usage. Other methods potentially

suitable for the inhibition of cyanobacterial blooms are direct grazing by phytoplanktivorous

fish (Jancula et al. 2008), biomanipulation (Lacerot et al. 2013) or a physical treatment (e.g.,

by ultrasonication, electroporation and cavitation) (Hao et al. 2004).

Cavitation is defined as the combined phenomena of the formation, growth and sub-

sequent collapse of microbubbles or cavities occurring over an extremely small interval of

time (milliseconds) and releasing large magnitudes of energy at the site of transformation

(Gogate 2011). Very high energy densities (energy released per unit volume) are obtained

locally, resulting in high pressures (in the range of 100–5,000 bar) and temperatures (in the

range of 1,000–10,000 K), and these effects are observed at millions of locations in the

reactor at one time (Suslick 1989). Although the effects of acoustic cavitation (ultrasound,

sonication) on bloom-forming cyanobacteria have been studied extensively (Hao et al.

2004; Joyce et al. 2010; Tang et al. 2004; Wu et al. 2012, 2011; Zhang et al. 2009),

knowledge about hydraulic cavitation still remains very limited (Xu et al. 2006).

In the present study, we examined the effects of hydrodynamic jet cavitation on the

survival, metabolic activity and ultrastructure of a bloom-forming cyanobacterium Micro-

cystis aeruginosa. Moreover, the impact of jet cavitation on a non-target green alga Chlorella

kessleri was investigated to compare the effects on cyanobacteria and green algae.

Materials and methods

Hydraulic jet cavitation device

An experimental circuit was built consisting of a water tank, centrifugal pump (Lowara,

rated output power 11 kW) controlled by a frequency convertor, a set of pressure

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transducers p1 and p2 (both BD sensor, 0–6 bar) and a flowmeter Q (Krohne). A con-

verging–diverging (CD) nozzle with inlet diameter 53 mm and minimum diameter of

30 mm in the throat section was manufactured from plexiglass. The nozzle was designed to

achieve a sufficient pressure drop due to cross-sectional area reduction and produce a

cavitating region filled with saturated vapor (i.e., hydrodynamic cavitation according to

Bernoulli principle is established). Since the vapor pressure is sensitive to the water

temperature and the amount of dissolved gas, a thermometer T (Sensit, type PT100) and

probe measuring dissolved oxygen content O2 (Oxymax) were incorporated into the

experimental circuit (the amount of dissolved oxygen was around 8.3 mg L-1 throughout

the measurements). The tests were carried out with a discharge of 18.4 L s-1; the pressure

difference in between both pressure probes was pulsating around a value of 350 kPa. The

static pressure in the throat reached the level of saturated vapor pressure (*3,500 Pa),

which was experimentally checked. Obviously, vigorous cavitation was also visually

observed in the whole nozzle beginning with the throat section (see Figs. 1, 2).

Test organisms

For the testing, laboratory cultures of cyanobacterium M. aeruginosa (isolated by E. Za-

pomelova from Vranov Reservoir in 2000) and C. kessleri (strain LARG/1) in ZBB growth

medium (1:1 mixture of medium Z (ZEHNDER in Staub 1960) and medium BB (Bristol

modified by Bold (1949) diluted to 50 % with distilled water)) were used. For the con-

firmation of the laboratory experiment and applicability in a real ecosystem, the natural

biomass of cyanobacterial water bloom from Brno Reservoir, Czech Republic (with M.

aeruginosa and Microcystis viridis domination), was further used in the experiments.

Design of the cavitation trials

The culture of organism tested (i.e., cyanobacterium or green alga) was kept in the storage

tank of the cavitation device in the amount of approximately 3 L. Based on the known flow

rate and the culture volume, we were able to detect the number of cycles which organisms

were exposed to. In the trials, cyanobacterium and alga were cavitated 0 (control), 1, 2, 4,

6, 12 and 18 times. The initial cell density in the experiments was established as a

Fig. 1 Scheme of hydraulic jet cavitation device

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compromise between relevance for a real ecosystem and the ability to measure the end

points in the laboratory by our instruments (flow cytometer, spectrophotometer and fluo-

rometer). At the beginning of the tests, cyanobacterial (M. aeruginosa) concentration was

approximately 50 lg L-1 (hygienic limit for bathing waters) of chlorophyll a and con-

centration of green alga (C. kessleri) was approximately 200 lg L-1 (relevant concen-

tration in eutrophic waters). According to flow cytometry, these chlorophyll

a concentrations corresponded to approx. 490,000 cells mL-1 (M. aeruginosa) and approx.

1,490,000 cells mL-1 (C. kessleri).

Determination of destruction of cyanobacterial gas vesicles

The effect of cavitation on the integrity of cyanobacterial (i.e., M. aeruginosa) gas vesicles

(vacuoles) was studied by flow cytometry. For this purpose, the side scatter parameter of

cyanobacterial cells was monitored. Since the intensity of the SSC signal reflects cell

granularity, damage to cyanobacterial gas vesicles results in a decrease in SSC (Lee et al.

2000). Histograms of SSC signal clearly showed that two populations of Microcystis cells

exist, both cells with a high SSC signal representing cells having intact gas vesicles and

low-SSC cells with damaged (ruptured) gas vesicles. Destruction of gas vesicles was also

checked by light microscopy (using an Olympus BX60 microscope).

Determination of cell counts immediately after the cavitation

Total cell counts were determined both in experimental organisms (i.e., cyanobacterium M.

aeruginosa and green alga C. kessleri) exposed to the cavitation treatment and in their non-

treated controls. This parameter was measured using a flow cytometer CyFlow ML (Partec,

Muenster, Germany) equipped with a blue excitation laser (488 nm/20 mW). Red chlo-

rophyll a autofluorescence emission (675/20 nm) was used as a trigger parameter enabling

accurate distinguishing of algal cells from the background. Concentrations of algal cells in

sample suspensions were investigated in a so-called true volumetric counting measurement

regime; thus, no reference particles (counting beads) should be used for flow cytometric

analysis.

Determination of cell membrane integrity

Membrane integrity was investigated using a fluorescent probe SYTOX Green as was

previously described by Regel et al. (2004) and Daly et al. (2007) with slight modifications

(Mikula et al. 2012). SYTOX Green is a specific green fluorescent dye which is unable to

cross non-damaged membrane of a healthy cell. Nevertheless, if the cell membrane is

Fig. 2 Details of the cavitation nozzle

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compromised (e.g., in chemically stressed cells), SYTOX Green may reach the cell and

bind to its nucleic acids. Stock solution of SYTOX Green (5 mM; Invitrogen, cat. number

S7020) was diluted by sterile ultrapure water to give the working solution of 50 lM.

Subsequently, 10 lL of working solution was added to 990 lL of sample into the flow

cytometric tube to obtain the final SYTOX concentration of 0.5 lM, and the sample was

stained for 7 min in the dark. Green fluorescence of cyanobacterial cells was detected using

a standard emission band-pass filter (527/30 nm). Heat-treated (80 �C, 10 min) cells were

used as a positive control.

Determination of metabolic (esterase) activity

Metabolic activity of cyanobacterial and algal cells was studied by flow cytometry using a

slightly modified version of Jochem’s protocol (Jochem 1999). The non-polar non-fluo-

rescent substrate fluorescein diacetate (FDA), due to its chemical structure, enters the cell,

where it is hydrolyzed by intracellular enzymes (mainly esterases) to fluorescein. This

fluorescent product emits green fluorescence, when excited by blue light. Moreover,

fluorescein remains trapped within the cell due to its polar character. Thus, the higher the

intensity of green fluorescence, the higher the metabolic activity of the cell. A stock

solution of FDA of 5 mg L-1 in dimethyl sulfoxide (DMSO) was stored at 4 �C. Before

measurement started, the stock solution was thawed and 10 lL was added to the Eppendorf

tube filled with 990 lL of sterilized ultrapure water. The prepared working solution

(50 lg L-1) was mixed carefully on the vortexer, and 33 lL was added to the sample into

the flow cytometric tube (with a final volume 1 mL). During analyses, a thawed stock

solution of FDA was kept on ice and the new batch of working solution was prepared for

each sample to prevent the possibility of substrate precipitation/degradation (Jochem

1999). Samples were incubated for 13 min in the dark, since our preliminary experiments

proved that these conditions are optimal for this kind of measurement. Cells were analyzed

using flow cytometry, and their green fluorescence intensity was detected using a 527/30-

nm band-pass emission filter.

Growth inhibition assays

Growth inhibition assays with both cyanobacterium (M. aeruginosa) and green alga C.

kessleri were performed immediately after the cavitation experiment. Untreated cyano-

bacterium and alga served as controls. The tests, based on ISO (1989), were performed in

transparent, 96-well microplates, each well of 250-ll volume, 3 replicates for each con-

centration and a control. Organisms in tests were incubated for 72 h at a temperature of

25 �C and light intensity of 100 lmol m-2 s-1 (6,000 lx) (Quantum sensor EMS 12,

Czech Republic) under fluorescent white cool lamps (PHILIPS). A 1:1 mixture of medium

Z (ZEHNDER in Staub 1960) and medium BB (Bristol modified by Bold (1949)) diluted to

50 % with distilled water was used as a growth medium. At the end of the test, the density

of algal cells was assessed using absorbance measurement performed with a microplate

reader (Sunrise, Tecan, Switzerland) at a wavelength of 680 nm.

Removal of natural biomass from the water column

To assess whether the hydraulic cavitation is able to remove natural cyanobacteria from the

water column, the following experiment was performed. Natural biomass with Microcystis

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sp. dominance was treated in the same design as laboratory cultures. Biomass was diluted

(by natural water from the same locality) to reach approximately 50 ll L-1 of chlorophyll

a. Cavitated biomass was sampled after 1, 2, 4, 6, 12 and 18 cycles into the 200-ml

beakers. After 24 h of standing, the surface layer (50 mL) and the bottom (50 mL) layer

were sampled carefully by pipette. The rest of the water (100 mL) served as a represen-

tative of the middle part of the water column in the beaker. The amount of the phyto-

plankton assemblage in every part of the water column (surface, middle and bottom) was

assessed by a fluorometer FluoroProbe (BBE Moldaenke, Kiel, Germany) equipped with a

cuvette adapter.

Data analysis

The experiments were repeated in three independent experiments, and each trial was

triplicated. ANOVA followed by a Tukey’s HSD post hoc test was conducted to analyze

differences between treated and untreated (controls) samples. Statistical analyses were

performed using statistical software Statistica 8 (StatSoft Inc., Tulsa, OK, USA), and

p \ 0.05 was considered to be significant.

Results and discussion

Determination of destruction of cyanobacterial gas vesicles

According to our data, the cavitation caused the degradation of 47.86–97.19 % (1–18

cavitation cycles used) of gas vesicles in M. aeruginosa. The histograms reflecting the

peaks of cells with low and high granularity are seen in Fig. 3. The lack of gas vesicles in

treated cyanobacterial cells was observed under the light microscope, as well (see Fig. 4).

No harm to cell membranes was detected under the microscope observation. This obser-

vation was confirmed also by flow cytometric counting before and after each cycle of the

cavitation process.

This finding is consistent with the results recently published by Zhang et al. (2009)

where the authors combined acoustic cavitation and coagulation to remove cyanobacterial

blooms. The main mechanism involved the destruction during ultrasonic irradiation of gas

vacuoles inside algae cells that acted as ‘‘nuclei’’ for acoustic cavitation and collapse

during the ‘‘bubble crush’’ period, resulting in the settlement of cyanobacteria (Zhang et al.

2009).

In a different study, Mahvi and Dehghani (2005) did trials with 150-s acoustic cavi-

tation (155-W input power, 42 kHz). The exposure effectively settled naturally growing

algae suspension. Sedimentation was caused by the disruption and collapse of gas

vacuoles.

Acoustic cavitation was successful in blue-green algae removal also in other studies as

Bott and Tianqing (2003), Zhang et al. (2006), Joyce and Mason (2008), Kotopoulis et al.

2009) and others.

Determination of cell counts and membrane integrity immediately after the cavitation

Although there was a slight decrease in cyanobacterial cell numbers after 18 cycles of

cavitation, the difference between the numbers before and after the cavitation was found to

be insignificant (ANOVA, p \ 0.05). Contrary to this, the amount of green algae increased

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from 1.49 9 106 cells mL-1 before the cavitation to 1.61 9 106 cells mL-1 immediately

after one cavitation cycle. This increasing trend was gradually progressing, and after 18

cavitation cycles, the amount of C. kessleri was found to be 1.74 9 106 cells mL-1. This

change in cell numbers was statistically significant (ANOVA, p \ 0.05). It is unlikely that

green alga grew during such a short period as one cycle of cavitation (approx. 10 s). We

Fig. 3 Histograms reflecting the peaks of M. aeruginosa cells with low side scatter (SSC) having a lowgranularity due to intact gas vesicles after 0 cycles (a), 1 cycle (b), 2 cycles (c) 4 cycles (d), 6 cycles (e), 12cycles (f) and 18 cycles (g)

Fig. 4 Light microscope image of cavitated (a) and uncavitated (b) Microcystis biomass

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believe that this is the result of disintegration of several small algal aggregates found in the

C. kessleri suspension.

Cell membrane integrity as an indicator of cell functionality was measured using a

fluorescent probe SYTOX Green. Membrane-compromised cells can be considered as

‘‘death,’’ since their internal structures become unprotected and they can be freely exposed to

the environment (Nebe-von-Caron et al. 2000). The resistance of such cells is limited, and

sooner or later, a cell undergoes the lysis. It was reported earlier that the chlorination of M.

aeruginosa cells may effectively disrupt the integrity of exposed cyanobacteria but also

causes a release of cyanobacterial toxins into the water environment which is even faster than

toxin degradation (Daly et al. 2007). From this perspective, the suitability of this method for

the water-bloom management seems to be questionable. Contrary to this, our results (see

Fig. 5) suggested that the cavitation did not cause any fundamental changes to cell mem-

brane integrity. The percentage of SYTOX negative cells (i.e., intact cells) was found to

range from 93.71 to 94.08 % of cells. The results of microscope observations, cell counts and

membrane integrity assessment are important for the detection of potential negative effects

on fauna and flora in aquatic ecosystems since the damage to cyanobacterial cells could lead

to the release of toxic cyanobacterial metabolites into the water. As we proved above, no

risks should arise as a consequence of hydraulic cavitation for bloom management.

Zhang et al. (2006) published time-dependent effects of acoustic cavitation (sonication)

on destruction of cyanobacterial cells. Whereas sonication for 1 and 5 s gave the highest

algal removal efficiency by gas vacuole collapse, longer exposure caused destruction of

algal structures themselves. Moreover, algal compartments spread into the water column

caused reduction in cell sedimentation. Similarly, destruction of cells was enhanced by

power above 64 W. In both cases, fragments of cells were well observable under micro-

scope, and thus, potential risks may be easily detected by this method.

Cell fragments were observed also in a study of Zhang et al. (2006). Cell destruction

was detected after 5 min of sonication (80 W, 80 kHz). Such undesirable treatment had an

impact on release of cyanobacterial toxins, microcystins. Microcystin (MC) concentration

in treated water increased up to 3.11 lg L-1 of MC.

Determination of metabolic (esterase) activity

To investigate the potential metabolic changes in cyanobacterial cells during the cavitation,

we examined the esterase activity of cells by using FDA as an esterase substrate. It was

Fig. 5 Membrane integrity of M. aeruginosa cells treated by hydraulic jet cavitation. The percentages ofboth SYTOX Green positive, i.e., membrane-compromised cells (white portions of columns) and SYTOXGreen negative, i.e., intact cells (gray portions of columns) after different numbers of cavitation cycles weredepicted. No significant differences were found between control and cavitation-treated groups

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found that the exposure of phytoplankton cells to various chemicals may affect their

esterase activity, most commonly in a negative manner (Hadjoudja et al. 2009; Jamers

et al. 2010; Mikula et al. 2012). Moreover, environmental factors such as a lack of nutrients

or low illumination intensity may decrease the metabolic activity of phytoplankton as well

(Brookes et al. 2000; Jochem 1999). Since temperature was previously found to influence

the metabolic activity of phytoplankton (Wu et al. 2008; Li et al. 2011), we kept the

temperature stable during our experiments, and when needed, we protected the tested

suspensions of cyanobacteria and green algae against warming up by cooling the cavitation

device during our experiments. Our results suggest that esterase activity was not influenced

by cavitation. The effects of cavitation on esterase activity are illustrated in Fig. 6.

Metabolic activity has been studied also by other authors. For example, Francko et al.

(1994) investigated the influence of nitrogen fixation by the cyanobacterium Anabaena

flos-aque. Anabaena reacted on cavitation by higher nitrogen fixation, and the frequency of

heterocysts was markedly higher in treated cultures over the duration of 5-day experi-

mental periods (Francko et al. 1994). Oxygen evolution and photoactivity decrease, on the

other hand, were studied by Zhang et al. (2006). According to this study, the cavitation

strongly damaged the antenna complexes and inhibited in total the photosynthesis of

M. aeruginosa.

Growth inhibition assays

Metabolic activity is closely connected to ability of cells to proliferate. In following trials,

we tried to detect the capability of phytoplankton species to grow in precisely defined

growth medium and external conditions after the cavitation. Our trials showed that

cyanobacterial growth was not influenced by hydraulic cavitation unlike the green alga

growth (see Fig. 7). This phenomenon may be explained by the ability of cavitation to

separate some aggregated Chlorella cells (see the first paragraph of ‘‘Results and discus-

sion’’) which are subsequently able to grow better in ZBB medium since their surface is in

greater contact with growth solution. The fact that higher cell numbers were observed for

alga and not for cyanobacterium after multiple cavitation cycles could be explained by the

higher number of cell aggregates in the culture of C. kessleri than in the culture of M.

aeruginosa. Theoretically, another reason could be the stimulation of green algae by

cavitation, but this is highly improbable especially when the higher cell numbers were

Fig. 6 Esterase activity of cyanobacterial (M. aeruginosa) cells after cavitation treatment. No significantdifferences were found between control and cavitation-treated groups

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observed immediately after the cavitation. The only paper dealing with the removal of M.

aeruginosa after the hydraulic cavitation seems to be the one from Xu et al. (2006).

However, the authors do not describe the degradation of gas vacuoles degradation. Xu et al.

(2006) describes the inhibition of growth where the inhibitive efficiency of 64.58 % in a

field water sample was detected. Although we did not observe such inhibitive effects on

cyanobacterium growth, we suppose (as well as the authors of the above-cited paper) that

such effects may be reached in strong dependency on the hydraulic characteristics of the

cavitation tube, pressure, number of cycles, cyanobacterial cell concentrations, etc. This

was confirmed by Hao et al. (2004) who found out that the growth inhibition mechanism

can be mainly attributed to the mechanical damage to the cell structures (caused by

ultrasonic cavitation), confirmed by light microscopy and differential interference

microscopy. On the other hand, if the ‘‘cell structures’’ are cell membranes, this could lead

to the release of toxins into the water. From this point of view, our cavitation device seems

to be more environmentally friendly, since the membrane integrity was not affected.

Removal of natural biomass from the water column

Since the theoretical assumptions for successful removal were accomplished (destruction

of gas vesicles, intact cell membrane, no harm to competitors such as green algae), we

decided to cavitate the natural biomass of Microcystis sp. to prove the efficiency of the

method under natural conditions. Trials with natural cyanobacterial biomass showed that

even one cavitation cycle is able to remove 66 % of cyanobacteria from the water surface

followed by 73 % (after 2 cycles), 83 % (after 4 cycles), 94 % (after 6 cycles), 97 % (after

12 cycles) and 99 % (after 18 cycles) of cyanobacteria. As Fig. 8 shows, this surface

removal was followed by accumulation of cyanobacteria without gas vesicles at the bottom

of testing beakers. The highest removal ratio of M. aeruginosa published in the literature so

far was 93.5 % by the sonication followed by coagulant application (Zhang et al. 2009). In

another study, Heng et al. (2009) acoustically cavitated (40 kHz, 60 W, 15 s) biomass of

M. aeruginosa. Cavitation improved algae coagulation removal by 12.4 % as compared

with direct coagulation.

Questions about possible application in natural conditions were discussed and also

answered in detail by Lee et al. (2002). The authors used cavitation for treatment of Lake

Senba. The water and sediment quality were monitored for 2 years, and the authors claim

Fig. 7 Absorbances of M. aeruginosa (gray columns) and C. kessleri (black columns) in control andcavitation-treated groups after 72 h of growth inhibition test. An asterisk indicates significant difference(p \ 0.05) in comparison with the control variant

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that cyanobacterial blooms can be controlled effectively in terms of chlorophyll a COD

and total phosphorus decrease.

Recently, Rajasekhar et al. (2012) published a review of acoustic cavitation to control

cyanobacterial blooms. The authors describe another result regarding direct removal of

algal biomass as well as inhibition of growth of cells. Moreover, the article discusses in

depth the effects on cyanobacterial metabolism and describes almost all sonication

parameters affecting the efficiency of the method.

Conclusions

Hydraulic cavitation seems to be a promising tool for the management of cyanobacterial

blooms as we presented in the paper. Until now, several authors have discovered effects of

acoustic cavitation (sonication) on cyanobacteria (Zhang et al. 2009; Tang et al. 2004; Hao

et al. 2004) and one team of authors the effects of hydraulic cavitation (Xu et al. 2006). We

believe that hydraulic cavitation seems to be more promising. Of course, the method has

weak points and some limitations for large-scale application like other measures used for

controlling of blue-greens. For example, the biomass at the bottom of the water body may

grow because its metabolic apparatus is apparently not damaged and the growth of cavi-

tated biomass was confirmed by our experiments. Moreover, the gas vesicles of living

cyanobacterial cells may be re-synthesized as published by Lee et al. (2000). On the other

hand, ultrasound and other physical and physicochemical methods deal with the same

problems, and despite this fact, they are used for correction of cyanobacterial blooms. The

cavitation is moreover provided through the separated (from surrounding water) cavitation

nozzle, and thus, aquatic vertebrates should not be affected as, for example, in the case of

the ultrasound method. Large-scale hydraulic cavitation can be used as an engine for boats

and thus combine recreation with cyanobacterial bloom management. Cavitation in whole-

scale application and subsequent precise monitoring should be done to verify the potential

of hydraulic cavitation against cyanobacterial water blooms as we presented in this study.

Acknowledgments We would like to thank Eliska Zapomelova for providing the laboratory strain of M.aeruginosa. The research was supported by long-term research development project no. RVO 67985939(Academy of Sciences of the Czech Republic). We gratefully knowledge the Czech Science Foundation for

Fig. 8 Removal of natural biomass from the water column in beaker trials. The bars represent percentage ofbiomass in the surface (white bars), in the middle of the column (light gray) and in the bottom (dark gray)after different numbers of cavitation cycles. An asterisk indicates significant difference (p \ 0.05) incomparison with the control variant

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support of the research under project No. 101/09/1715 ‘‘Cavitating vortical structures induced by rotatingliquid’’ as well.

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