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LIMNOLOGY
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OCEANOGRAPHY: METHODS
Limnol. Oceanogr.: Methods 6, 2008, 513–522
© 2008, by the American Society of Limnology and Oceanography, Inc.
In situ bacterial mitigation of the toxic cyanobacterium
Microcystis aeruginosa: implications for biological bloom control
Baik-Ho Kim1*, Miao Sang2,3, Soon-Jin Hwang1, and Myung-Soo Han2*
1
Department of Environmental Science, Konkuk University, Seoul 143-701, Republic of Korea
Department of Life Science, Hanyang University, Seoul 133-791, Republic of Korea
3
Department of Environmental Microbiology, Harbin Institute of Technology, Harbin 150-001, China
2
Abstract
The algicidal bacterium Xanthobacter autotrophicus HYS0201-SM02 (SM02) was isolated from the surface water
of a eutrophic lake (Lake Daechung, Korea). In vivo and in situ experiments showed that SM02 had algicidal
activity against both a cultured strain and natural colonial morphs of the toxic cyanobacterium, Microcystis
aeruginosa. Both the SM02 bacteria and its culture filtrate showed anti-algal activity against M. aeruginosa, indicating that an algicidal substance was released from SM02. The threshold concentration of SM02 for maximal
algicidal activity against a natural bloom of M. aeruginosa was 107 CFU/mL. In situ co-culture of SM02 and M.
aeruginosa showed that SM02 did not benefit from the massive decay of M. aeruginosa. In fact, repeated inoculations with a low concentration of SM02 were required for optimal algicidal activity, suggesting that water
quality worsened during co-culture (i.e., nutrients and microcystin-LR concentration increased). These results
suggest a role for the algicidal bacterium X. autotrophicus SM02 in biorestoration but probably not in treating
outdoor Microcystis blooms. When developing a biological agent to control M. aeruginosa blooms in the field, it
will be important to screen for specific agents with low threshold concentrations and high algicidal activity.
Introduction
Direct application of chemicals to control algal blooms can
harm aquatic ecosystems by killing beneficial organisms such
as plankton and fish (McGuire et al. 1984; Reynolds 1984;
Reyssac and Pletikosic 1990). Therefore, many researchers
have taken pan-ecological and environmental approaches to
lake conservation by investigating the potential for biological
control of cyanobacterial blooms, mainly using Microcystis
aeruginosa as a model system (Redhead and Wright 1978;
Brabrand et al. 1983; Manage et al. 2000; Choi et al. 2005; Kim
et al. 2007). So far, most reports have been from laboratory
studies, and the biological control of algal blooms in situ has
not been adequately assessed.
There is ongoing debate, particularly among Japanese
researchers, regarding the effect of heterotrophic bacteria on
the growth dynamics of harmful freshwater and marine algae
(Yamamoto et al. 1998; Manage et al. 2000; Kodani et al.
2002). Although numerous laboratory studies have demonstrated anti-algal effects of some bacteria on marine and freshwater algae species (Daft et al. 1975; Sigee et al. 1999; Lee et al.
2000; Mayali and Azam 2004; Kim et al. 2007), the use of M.
aeruginosa as an agent of biological control in situ, that is,
under natural conditions or in bloom water, is complicated.
Choi et al. (2005) demonstrated that the Microcystis-killing
bacterium Streptomyces did not continue to grow as the number of algae declined and that bacterial growth required the
As a result of eutrophication, cyanobacterial blooms have
become common in lakes and reservoirs worldwide (EkmanEkebom et al. 1992). Such blooms result in foul odors,
decreased aesthetic value, deteriorated water quality, and water
deoxygenation (Falconer 1999; Oberholster et al. 2004).
Cyanobacterial blooms can also lead to the production of
microcystins (MCs) that can be toxic to aquatic organisms
including fish, birds, wild animals, livestock, waterfowl, as well
as to humans (Carmichael et al. 1975; Codd et al. 1989; Jacquet
et al. 2004; Ernst et al. 2006; Palikova et al. 2007). MCs are also
associated with allergies, irritant reactions, gastroenteritis, liver
disease, and tumors in humans (An and Carmichael 1994; Bell
and Codd 1994; Dawson 1998; Almeida et al. 2006).
*Corresponding authors: E-mail: [email protected] (B.H.K),
[email protected] (M.S.H)
Acknowledgments
We are grateful to Dr. N. Takamura and J. P. Gaur for valuable comments on earlier versions of the manuscript. S.W. Jung and J.K. Seo provided field and technical assistance. Han-River Environmental Research
Laboratory (HERL) provided a small pond for the field study. This work was
supported by a Korea Research Foundation Grant (KRF-2004-C00018).
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developed on the lawn. Bacteria associated with these zones
were isolated by serial streaking onto nutrient agar (NA) plates.
The isolated anti-algal bacteria were cultured in nutrient broth
(NB) in the dark at the optimal temperature of 40°C, with gentle orbital shaking (120 rpm). Bacterial biomass was determined by dry weight (APHA 1998). The isolates were maintained axenically in the dark on NA plates and cryopreserved
at –76°C in NB containing 20% glycerol.
Identification of algicidal bacteria—To identify the algicidal
bacterial isolates, bacterial chromosomal DNA was isolated
according to the method described by Choi et al. (2005), and
molecular identification was performed as in Wellinghausen
et al. (2005). Briefly, 16S rDNA was amplified using primers
27F, 5′-GAGTTTGATCATGGCTCAG-3′ and 1492R, 5′-GGTTACCTTGTTACGACTT-3′, in a 50 μL reaction volume containing 20 ng of template DNA, 1× PCR buffer (Sigma), 5 mM
MgCl2, 10 mM dNTPs, 10 pM of each primer, and 2.5 units of
Taq DNA polymerase. The PCR consisted of 35 cycles of denaturation for 1 min at 94°C, annealing for 2.5 min at 55°C,
extension for 2.5 min at 72°C, and a final elongation step of 7
min at 72°C in a DNA thermal cycler (Genetic analyzer 377;
Perkin-Elmer). The PCR products were purified with the QIA
quick PCR Purification Kit (Qiagen) and sequenced by automated DNA sequencing (Bionex). The resulting 16S rDNA
sequences were aligned using CLUSTAL W software (Thompson et al. 1994), and the distance matrices were calculated
using the DNADIST program within the PHYLIP package
(http://evolution.genetics.washington.edu/phylip.html). A
phylogenetic tree was constructed by the neighbor-joining
method based on the calculated distance matrix. Identification of the HYS0201-SM02 isolate (SM02) was determined by
full-length 16S rDNA sequence comparison with sequences in
the DDMJ/EMBL/Genbank database. The sequence of the
SM02 isolate was similar to the sequences of various strains of
Xanthobacter and was identical to those of X. autotrophicus
T101 (GenBank accession no. U62887), T102 (U62888), and
7c (X94201) (Fig. 1). The sequences of the SM02 isolate were
deposited in GenBank under the accession number EU982303.
These aerobic gram-negative bacteria are known to decompose
1,2-dichloroethane, a toxic, halogenated compound commonly
used in the production of vinyl chloride (Baptista et al. 2006) and
have been reported to produce the acetone-using enzyme, acetone carboxylase (Sluis and Ensign 1997; Sluis et al. 2002).
Optimal growth of algicidal bacteria—To determine the optimal growth conditions for SM02, the isolate was grown with
gentle shaking in NB (pH 7) at 25°C, 30°C, 35°C, 40°C, 45°C,
and 50°C. It was also grown in NB at 40°C at various pHs (5,
6, 7, 8, 9, and 10). Bacterial growth was quantified by 4’,6diadimino-2-phenylindole (DAPI) staining (Porter and Feig
1980). Briefly, bacteria fixed with 2% glutaraldehyde (final
concentration) were filtered onto black polycarbonate filters
(0.6-μm pore diameter) and counted at 1000× magnification
using an epifluorescence microscope (Karl Zeiss). The optimal
growth conditions determined by these experiments were
addition of nutrients (e.g., casitone), because the target algae
were not a sufficient source of food. The need for added nutrients when applying bacteria for algal bloom control in
eutrophic waters is an important constraint. Sigee et al. (1999)
and Mayali and Azam (2004) also reported that the conditions
optimal for growth of algicidal bacteria can differ from those
used in experiments and those used for algal growth control.
Typically, conditions optimal for the growth of the target alga
are used in co-culture experiments involving bacteria, potentially resulting in low algicidal activity.
Evidence that algicidal bacteria can control blooms of M.
aeruginosa is based on reports of increased bacterial abundance
correlating with the termination of cyanobacterial blooms (Shilo
1967; Daft et al. 1975; Rashidan and Bird 2001). Investigation
into the algicidal mechanisms of bacteria has shown that some
bacteria actually increase the concentration of dissolved MCs via
lysis of Microcystis cells (Choi et al. 2005; Kim et al. 2007). The
mechanism of MC-mediated destruction of Microcystis and the
relationship between MC and algicidal bacteria remain unclear.
Moreover, there are no reports about using algicidal bacteria to
control colonial morphs of M. aeruginosa in situ.
In this report, we describe bacteria-mediated control of M.
aeruginosa. Specifically, we isolated and characterized a Microcystis-lytic bacterium (Xanthobacter autotrophicus) with host
specificity. The algicidal activity of X. autotrophicus was
assessed at different bacterial densities as well as at different
bacterial and algal growth phases. Finally, we performed an in
situ test to determine whether the algicidal activity (AA) persisted in suboptimal bacterial growth conditions within a natural phytoplankton community.
Materials and procedures
Study of algae and algicidal bacterial isolation—The toxic
cyanobacteria M. aeruginosa NIER-100001 were obtained from
The National Institute for Environmental Research (NIER,
Korea) and cultured in modified Cary-Blair (CB) medium
(Watanabe and Hiroki 1997) on an orbital shaker (120 rpm) at
25°C with a 12 h:12 h light:dark cycle (light intensity 50 μE m–2
s–1). The M. aeruginosa cells used in this study were spherical or
ovoid, 3.7 to 5.7 μm in diameter, and 27 to 97 ìm3 in volume.
Bacteria with algicidal activity against M. aeruginosa NIER100001 were isolated from an algal lawn or through the use of
a modified soft agar overlay method (Sakata et al. 1991). Surface water samples were collected from Lake Daechung, Korea,
during an algal bloom on June 12, 2004, and were then filtered
through 0.8-μm nucleopore membrane filters. Cultures of M.
aeruginosa N-100001 were grown in CB medium for 7 d and
harvested by centrifugation at 3000×g for 20 min. The cell pellet was mixed with molten CB soft agar medium and poured
into a plastic Petri dish (87-mm diameter). A cyanobacterial
lawn developed after 2 to 3 d, and 200-μL water samples were
spread onto the lawn. After incubation for 5 to 10 d, the plates
were examined for the presence of clear zones (indicating inhibition of cyanobacterial growth) around bacterial colonies that
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100 mL flasks containing 50 mL of mid-exponential phase
M. aeruginosa (13.57 × 105 cells mL–1, cultured for 13 d). The
M. aeruginosa cells were counted daily, as described below.
Anti-algal activity of SM02 against other algae—We tested the
AA of SM02 in liquid cultures of cyanobacteria Anabaena cylindrica (NIES-19; NIES; National Institute for Environmental
Studies), A. flos-aquae (KCTC-AG10026; KCTC; Korean Collection for Type Cultures, KRIBB, Daejeon, Korea), and M. aeruginosa (strains NIES-44, NIES-101, and NIES-298), and the
diatoms Aulacoseira granulata (CCAP 1002/1; CCAP; Culture
Collection of Algae and Protozoa), Cyclotella meneghiniana
(previously isolated by Han, M.S.), and Stephanodiscus
hantzschii (UTCC267; UTCC; University of Toronto Culture
Collection of Algae and Cyanobacteria).
Briefly, SM02 was cultured in NB in the dark (24 h, 40°C,
orbital shaking at 150 rpm). The bacteria were harvested by
centrifugation at 14,000×g for 20 min, and inoculated to the
cultures of each algal species, while the final bacterial density
was adjusted to 1×108 cells mL–1 for all algal species. Algae
samples were collected from exponential-phase cultures. The
cyanobacteria were cultured in 300-mL flasks containing 100
mL sterilized CB medium, as described previously (25°C, light
intensity 50 μE m–2 s–1, 12 h:12 h light:dark cycle, orbital shaking at 120 rpm). The three types of diatoms were cultured at
20°C in 300-mL flasks containing 100 mL sterilized diatom
medium (DM), pH 6.9 (Beakes et al. 1988). The light intensity,
light:dark cycle, and shaking conditions were as for cyanobacteria. The AA of SM02 was calculated using the same equation
as for the cyanobacteria.
In situ anti-algal activity of SM02 on natural Microcystis bloom
water—In 1981, we constructed a small artificial concrete
pond (ca. 300 m3) directly connected to the lower part of the
Han River (Paltang reservoir, Korea) for the purpose of field
experiments. Since its construction, the study pond has developed Microcystis blooms every year between April and November. The cyanobacterial blooms are composed of over 90% M.
aeruginosa and ~10% other phytoplankton species.
To assess the AA of SM02 in natural Microcystis bloom
water, we conducted an in situ experiment over 11 d (June 14
to June 24, 2004) in 12 cone-shaped PVC tanks. The PVC
tanks (0.8 m high; total volume 120 L) contained 100 L water
supporting a natural Microcystis bloom. The tanks were treated
with SM02 as follows: 1) no added bacteria (Control); high
concentration of bacteria (HD, 1 × 108 CFU/mL); midconcentration of bacteria (MD, 5 × 107 CFU/mL); low concentration
of bacteria (LD, 1 × 107 CFU/mL), and a bi-daily low-concentration (1 × 107 CFU/mL) inoculation performed five times
during the study (BLD, final added concentration, 5 × 107
CFU/mL, the same as for MD). All experiments were performed in triplicate.
The SM02 bacteria, prepared as described below, were
sprayed onto the water surface of each tank on the fifth day of
the 11-day experiment. Axenic cultures of SM02 were maintained in the dark on NA plates (1.5% agar; pH 7) at 20°C, or
Fig. 1. Phylogenetic tree based on 16S rDNA sequences, showing the
relative positions of the Microcystis-killing bacterium HYS0201-SM02, the
type strains of some Xanthobacter species, and the type strain of Xanthobacter autotrophicus. Scale bar represents 0.01 substitutions per
nucleotide position.
used for all subsequent bacterial cultures.
Anti-algal activity of SM02 on M. aeruginosa—Two methods, a
paper disc test and a liquid culture test, were used to assess the
anti-algal activity of the bacterial isolates and to select the isolate most effective against M. aeruginosa. The paper disc method
involved growing the isolates in NB medium at 40°C with shaking for 2 d. Discs of Whatman GF/F filter paper (pore size = 0.7
μm; 5-mm diameter) were soaked with 100 μL of each cultured
bacterial isolate, placed on lawns of M. aeruginosa, and incubated for 5 d. Anti-algal activity was recorded as the diameter of
the clear zone that formed in the algal lawn. For the liquid culture test, a bacterial culture was washed with algal medium and
then inoculated into a 50-mL test tube containing 25 mL of M.
aeruginosa N-100001 culture. The co-culture was incubated for 5
d. Each day, bacterial growth was determined by the DAPI
method, and algal growth was measured by direct counting
using an inverted microscope (Utermöhl 1958).
To assess the effect of SM02 on M. aeruginosa during different phases of algal and bacterial growth, a culture of
SM02 was prepared as described above, adjusted to an optical density of 1.8 at 660 nm, and 2.5 mL inoculated into
100 mL flasks containing 50 mL of lag, exponential, or stationary phase M. aeruginosa cells (2.3, 20.7, and 22.7 × 105
cells mL–1, respectively). We also measured the AA of SM02
at different growth phases (lag, exponential, and stationary phase) on M. aeruginosa in exponential phase. Aliquots
of SM02 were incubated for 3 h (lag phase), 18 h (exponential phase), and 36 h (stationary phase) in 100 mL
flasks containing 50 mL of NB. The cells were harvested by
centrifugation at 18,000×g for 20 min, the cell concentration was adjusted to 1.8 at 660 nm, and 2.5 mL (approximately 1 × 108 cells mL–1) of culture was inoculated into
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each sample, filtered with glass filters (25 mm diameter,
Whatman GF/C), and extracted with 10 mL of 90% methanol
for 24 h at room temperature. Each methanol-extracted sample (10 to 50 μL) was neutralized or diluted with distilled water
(90 to 50 μL). The samples or standards were mixed with an
appropriate dilution of M8H5 MAb and dispensed in a 96-well
microtiter plate (Coaster) pre-coated with an MC-bovine
serum albumin conjugate. The plates were washed, and the
bound MAb was detected with horseradish peroxidase-labeled
goat anti-mouse IgG (TAGO 4550) and its substrate (0.1
mg/mL of 3,3′,5,5′-tetramethylbenzidine, 0.05% H2O2 in 0.1
M acetate buffer, pH 5.0). Finally, absorbance was measured at
450 nm. The MC concentration was determined from a standard competitive curve with a detection limit of 50 pg/mL. For
the d-MC, small quantity (50-100 μL) of above 2-L filtered
water was used directly to measurement without methanol
extraction and water dilution. The next step was the same that
of c-MC measurement.
Data analysis—The AA of SM02 on cultured or natural
Microcystis blooms and/or other algae was calculated using the
equation AA (%) = (1–T/C) × 100, where T and C are the cell
densities of Microcystis in the presence or absence of SM02,
respectively. Differences in cell densities between treated and
control cultures were analyzed by analysis of variance
(ANOVA), and data were compared using linear contrasts. Values of P < 0.05 were considered statistically significant. Statistical analyses were performed using the SPSS package (SPSS
Inc. 1989-2003).
cryopreserved at –76°C in NB medium (Difco, South Royal)
containing 20% glycerol. Prior to the experiment, frozen bacteria were inoculated into 300 mL fresh NB medium and incubated at 25°C with shaking (120 rpm). After 48 h, 2.7 L NB
medium was added, and incubation was continued for 60 h.
The bacterial cells were then harvested by centrifugation at
18,000×g for 20 min and washed twice with distilled water.
The SM02 bacteria were introduced to the experimental tanks
within 24 h of harvesting.
Phytoplankton community and water chemistry—Before taking
water samples from the tanks, the physiochemical parameters
(water temperature, dissolved oxygen, conductivity, turbidity,
and pH) in each tank were measured at a depth of 50 cm using
a YSI portable meter (YSI-61). For measuring nutrient concentrations and phytoplankton densities and/or biomass, water
samples were collected from a depth of 50 cm in each tank,
transferred to a 10-L plastic bucket, and mixed. To enumerate
the phytoplankton, 250 mL subsamples were fixed with
Lugol’s solution (1% final concentration) in polypropylene
bottles and stored in the dark at 4°C until analysis. A 1-mL
sample of fixed phytoplankton was placed in a SedgwickRafter Counting Chamber (PhycoTech Inc.) and observed by
light microscopy (Olympus microscope, Tokyo, Japan) at 4001000× magnification. Phytoplankton were enumerated based
on counts of 200-300 cells per sample and identified at the
genus or species level. The total biomass (carbon content) of
phytoplankton was calculated using a conversion factor and
biovolume values (Mullin et al. 1966; Strathmann 1967).
For inorganic nutrient analysis, water samples were filtered
(Whatman GF/F; Whatman International) and stored frozen
(–10°C) until analysis. Ammonium (NH4), nitrate nitrogen
(NO3), and soluble reactive phosphorus (PO4) were measured
with an automatic analyzer (TRAACS800, Bran-Luebbe) after
pretreatment with ascorbic acid, indophenol blue, and cadmium reduction (APHA 1998). The silicone concentration was
measured by inductively coupled plasma-atomic emission
spectrometry (ICAP 61E Trace; Thermo Jarreell-Ash). To measure the concentration of chlorophyll a (Chl a), 100-mL subsamples were filtered (Whatman GF/F) and extracted
overnight with 90% acetone at 4°C. The fluorescence of each
extract was measured using a spectrophotometer calibrated to
Chl a standards (Sigma-Aldrich Co.).
Microcystin-LR measurements—The concentration of cellular
(c-MC) and dissolved microcystin-LR (d-MC) was quantified
from each sample by enzyme-linked immunosorbent assay
(ELISA; Nagata et al. 1997). The total concentration (t-MC)
was calculated as the sum of the c-MC and d-MC values.
Because the anti-microcystin monoclonal antibody (MAb)
M8H5 used in the ELISA assay reacts equally with all major
microcystin derivatives (i.e., microcystin-LR, microcystin-RR,
and microcystin-YR), we used 250 μg of the most widely studied microcystin, microcystin-LR (C49H74N10O12, MW = 995.17,
Wako) as the ELISA standard. Briefly, to quantify the concentrations of c-MC, 2-L samples of the water were taken from
Assessment
Algicidal activity of SM02 bacteria on toxic Microcystis aeruginosa—Optimal growth of the SM02 isolate occurred at 40°C
and pH 7, and the isolate was characterized by rod-shaped
cells, yellowish colonies, gram-negative staining, oxidase-positivity, and growth on D-maltose (data not shown).
We tested the AA of SM02 against the toxic cyanobacterium
M. aeruginosa NIER100001 in both the paper-disc and liquidculture tests. SM02 was co-incubated with M. aeruginosa for 5 d.
When the starting concentration of SM02 was 108 cells mL–1,
the algal cell density after 5 d was decreased by approximately
95.6% relative to the uninoculated control; that is, the algal cell
density was only 4.6% of the cell density in the control culture.
When the starting concentration of SM02 was 107 cells mL–1,
the algal cell density after 5 d was only 33.9% of the uninoculated control (Fig. 2A). In contrast, inoculation of SM02 at concentrations below 106 cells mL–1 did not effectively inhibit M.
aeruginosa (ANOVA, P > 0.5) as compared with the control.
There were significant differences between the AA of bacterial cell culture at 108 cells mL–1 and the AA of the corresponding culture filtrate (ANOVA, P < 0.0001). Interestingly, the filtrate from SM02 cultures at 107 cells mL–1 effectively
suppressed M. aeruginosa cell growth (to 51.5% of the control
after 5 d; Fig. 2B), exceeding the suppression by SM02 cell culture at 107 cells mL–1 and indicating that the bacteria secreted
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Fig. 3. Anti-algal effects of Xanthobacter autotrophicus HYS0201-SM02
on Microcystis aeruginosa N-10001 (A) at different bacterial growth
phases (lag, exponential, and stationary) against exponential phase M.
aeruginosa N-1000, and (B) at different algal growth phases (lag, exponential, and stationary). The arrow indicates the bacterial inoculation
time. Each value is the mean ±SD of three replicates.
Fig. 2. Anti-algal effects of Xanthobacter autotrophicus HYS0201-SM02
for exponential phase Microcystis aeruginosa N-10001 (A) at different bacterial cell concentrations, and (B) by bacterial cells and the corresponding
culture filtrate. The arrow indicates the bacterial inoculation time. Each
value is the mean ±SD of three replicates.
meneghiniana (2.7%). Therefore, SM02 effectively suppressed
the growth of toxic M. aeruginosa strains but had little effect
on the growth of diatoms or some other cyanobacteria,
including a different strain of M. aeruginosa, implying interand intra-specific host susceptibility.
In situ algicidal activity of SM02 on natural Microcystis water—
In situ, the biomass of M. aeruginosa was significantly reduced
(to approximately 73.9% of control) by the addition of a high
concentration (HD) of SM02 and by bi-daily additions of low
concentration (BLD) of SM02 (to approximately 54.7% of control; ANOVA, P < 0.001 for both treatments). Addition of a low
concentration (LD) of SM02 enhanced M. aeruginosa cell
growth significantly, to approximately 121.4% of control
(ANOVA, P < 0.001), while treatment with the middle concentration (MD) of SM02 did not have a significant effect
(ANOVA, P > 0.5; Fig. 4).
The effects of co-culturing SM02 at various concentrations
with other phytoplankton species varied (Fig. 4). The carbon
biomass of some phytoplankton increased after bacterial
an algicidal substance into the media. Notably, the bacterial
biomass did not change substantially after inoculation, but
fluctuated over the 7 d of incubation. Inoculates of SM02 in
the exponential and/or stationary growth phase had greater
AA as compared to cells in lag phase (Fig. 3A). In contrast,
exponential phase SM02 effectively suppressed M. aeruginosa
in all growth phases (Fig. 3B). Therefore, active SM02 at a density greater than 107 cells mL–1 controlled Microcystis growth in
all culture stages tested.
Algicidal activity of SM02 on other phytoplankton species—
During 5 d of co-cultivation, SM02 strongly inhibited two M.
aeruginosa strains, NIER100001 (97.5% growth inhibition) and
NIES101 (90.2%), and moderately inhibited NIES298 (46.6%;
Table 1). In contrast, SM02 had little effect on Anabaena cylindrica (31.7% growth inhibition), M. aeruginosa NIES44
(26.0%), Aulacoseira granulata (23.5%), Stephanodiscus
hantzschii (16.4%), Anabaena flos-aquae (8.8%), or Cyclotella
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Table 1. Antialgal activity (AA) of the isolate HYS0201-SM02 against various phytoplankton species measured by direct counting (or
Chl a concentration for Anabaena species) after 5-d co-culture.
Host strain
AA*
Anabaena cylindrica Lemmermann NIES19†
Anabaena flos-aquae (Lyngb.) Brebisson KCTCAG10026‡
Aulacoseira granulata (Ehr.) Simonsen CCAP1002/1§
Cyclotella meneghiniana Kützing HYK0210-CM01#
Microcystis aeruginosa (Kütz.) Lemmermann NIES44
Microcystis aeruginosa (Kütz.) Lemmermann NIES101
Microcystis aeruginosa (Kütz.) Lemmermann NIES298
Microcystis aeruginosa (Kütz.) Lemmermann NIER100001**
Stephanodiscus hantzschii Grunow UTCC267††
31.7 (25.2–42.4)
8.8 (6.6–11.4)
23.5 (18.2–27.0)
2.7 (–0.8–4.4)
26.0 (24.5–28.3)
90.2 (88.1–92.6)
46.6 (41.4–53.7)
97.5 (95.3–99.6)
16.4 (15.7–18.2)
All algal strains were cultured with gentle shaking at a light intensity of 50 μE m–2 s–1on a 12 h:12 h light:dark cycle. AA is presented as the mean (range)
of triplicate experiments.
*
AA (%) = (1 – T/C) × 100, where T and C are the algal cell densities in the presence and absence of HYS0201-SM02, respectively.
†
NIES; National Institute for Environmental Studies, Tsukuba, Japan.
†
KCTC; Korean Collection for Type Cultures, KRIBB, Daejeon, Korea.
§
CCAP; Culture Collection of Algae and Protozoa, Scotland, UK.
#
Strains were isolated by capillary method and deposited at Hanyang University, Seoul, Korea
**
NIER; National Institute of Environmental Research, Incheon, Korea.
††
UTCC; University of Toronto Culture Collection of Algae and Cyanobacteria, Toronto, ON, Canada.
inoculation: Chroococcus turgidus (approximately 8.5–10.4-fold
greater growth than control, P < 0.001 for all concentrations
tested), Scenedesmus quadricauda (approximately 21% greater
than control, P < 0.01 for only the LD treatment), and
Ankistrodesmus falcatus (approximately 17% greater than control, P < 0.001 for only the BLD treatment). Interestingly, the
carbon biomass of the green alga Coelastrum sphaericum
decreased in the presence of all concentrations of SM02. In particular, the BLD treatment suppressed algal biomass by approximately 92%. Thus, suppression of Coelastrum sphaericum by
SM02 was greater than the suppression of natural M. aeruginosa,
suggesting algicidal potential for SM02 beyond Microcystis.
All physicochemical water quality parameters tested
changed in the presence of SM02, including decreased dissolved oxygen, increased nutrients (N, P, and Si), and
increased turbidity (data not shown). As expected, there was a
significant increase in d-MC (ANOVA, P < 0.001 for HD, MD,
and BLD) and a significant decrease in c-MC concentration
(ANOVA, P < 0.0001 for BLD, P < 0.001 for HD and MD) after
5 d of co-culture with SM02 (Fig. 5), possibly due to algal cell
lysis or degradation. Interestingly, the concentration of Chl a
significantly increased in parallel with decreasing concentrations of M. aeruginosa after bacterial inoculation (ANOVA, P <
0.0001 for HD and MD, P < 0.001 for LD and BLD), suggesting
a phytoplankton succession (e.g., Microcystis to Chroococcus in
cyanobacteria; Coelastrum to Ankistrodesmus in green algae).
inhibition (> 90% growth inhibition) evident only in two
M. aeruginosa strains (NIES-101 and NIER-100001), and moderate or low inhibition evident only in M. aeruginosa strains NIES298 and NIES-44. SM02 also showed little inhibitory activity
against the toxic cyanobacterium Anabaena flos-aquae and the
small centric diatoms Stephanodiscus hantzschii and Cyclotella
meneghiniana. It is generally accepted that these intra- and interspecific differences susceptibility of alga to algicidal bacteria are
due to physiological differences among algal strains (Manage et
al. 2000; Walker and Higginbotham 2000; Yasuno et al. 2000),
bacterial density, and bacterial growth rates (Mayali and Azam
2004; Choi et al. 2005). Our results suggest that SM02 not be
effective against all species of M. aeruginosa. Therefore, the goal
of isolating bacteria that can mitigate the harmful effects of M.
aeruginosa remains elusive and vitally important.
In the in situ experiments with the Microcystis bloom water,
two concentrations of SM02 (HD and BLD) showed AA against
M. aeruginosa. Although SM02 (LD) significantly increased the
cell density of M. aeruginosa (P < .01), repeated LD treatments
(i.e., BLD treatment) was more effective in reducing M. aeruginosa than MD treatment of SM02. Interestingly, the total inoculated bacterial density and the corresponding AA from five
inoculations of SM02 LD were less than those of a single HD
treatment. Taken together, these results indicate that for optimal AA against a natural bloom of M. aeruginosa in situ, the
concentration of SM02 must be at least 107 CFU/mL. However,
it remains to be seen whether repeated treatment with SM02,
resulting in a higher total bacterial density than a single HD
treatment, will be effective against M. aeruginosa. In particular,
all bacterial concentrations tested strongly increased the biomass of cyanobacterium Chroococcus turgidus (approximately
8.5~11.5 fold-higher-growth) as compared with the control.
Discussion
We found that the bacterium X. autotrophicus SM02
strongly suppressed the growth of M. aeruginosa NIER-10001
in the laboratory. However, there was distinct variability in
algal susceptibility to the AA of SM02 (Table 1), with strong
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Bacterial mitigation of M. aeruginosa
Fig. 4. Phytoplankton biomass before and after addition of Xanthobacter autotrophicus HYS0201-SM02 to an isolated natural bloom of Microcystis aeruginosa. Control, no bacteria added into the tank; HD, high concentration of bacteria (1 × 108 CFU/mL); MD, mid-level concentration (5
× 107 CFU/mL); LD, low concentration (1 × 107 CFU/mL); and BLD, bidaily low-concentration (1 × 107 CFU/mL) inoculation performed five
times during the study (a total of 5 × 107 CFU/mL, equivalent to MD).
Fig. 5. Concentrations of Chl a, cellular (c-MC), and dissolved microcystin-LR (d-MC) before and after addition of Xanthobacter autotrophicus
HYS0201-SM02 into enclosed waters containing a natural bloom of
Microcystis aeruginosa. Values are means of triplicate experiments for the
five day periods 14–18 June (PRE) and 19–24 June 2005 (POST).
Moreover, BLD treatment increased the biomasses of green
algae such as Ankistrodesmus and Scenedesmus, while one-time
inoculations with other concentrations did not. We propose
that repeated treatment with algicidal bacteria, regardless of the
amount added, decreases the water quality due to the increase
in organic materials such as bacteria and medium debris.
During co-cultivation of SM02 and M. aeruginosa, the bacterial biomass fluctuated but did not increase substantially.
Thus, the bacterial population did not benefit from the massive decay of the M. aeruginosa population. The bacterial diversity, as determined by DGGE and FISH assay, did not change
when the addition of the filtered Microcystis bloom water to
the natural water (Kim, unpublished data). This phenomenon
could be related to the effect of the growth conditions on bacterial behavior or to the effect of toxic metabolites produced
by M. aeruginosa. As in a previous study using Streptomyces
(Choi et al. 2005), SM02 grew optimally at pH 7 and 40°C,
conditions different from those optimal for the cyanobacterium and different from those used during co-cultivation.
In addition, the concentration of dissolved d-MC gradually
increased relative to c-MC following SM02 inoculation (Fig.
5). Kim et al. (2007) demonstrated that the release of d-MC
from Microcystis following cell lysis by algicidal bacteria could
suppress growth of the anti-algal bacteria. Although a detoxification role for microcystin-degrading bacteria residing in the
outer mucilage of Microcystis has been proposed (Rashidan and
Bird 2001; Maruyama et al. 2003), the bacteriolytic activity of
M. aeruginosa via MC derivatives has not been tested in situ.
Therefore, we hypothesize that the low growth or biomass of
X. autotrophicus SM02 was due to unfavorable growth conditions (e.g., water temperature that was too low or too high).
Further studies are necessary to better understand the mechanisms of bacteria-mediated inhibition of M. aeruginosa. In particular, toxin production and bacteriolytic or alga-lytic
processes (e.g., Kitaguchi et al. 2001) following bacterial
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inoculation remain to be elucidated.
Since the initial report by Shilo in 1967, many different
Microcystis-killing bacteria have been isolated, including Streptomycetes phaeofaciens (Yamamoto et al. 1998), Alcaligenes denitrificans (Manage et al. 2000), Pseudomonas sp. (Kodani et al.
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aeruginosa) with the goal of biorestoration, with the exception
of a recent study by Takamura et al. (2004) involving the
direct spraying of the algicidal amino acid, L-lysine. The difficulties and limitations of in situ experiments experienced by
most researchers are most likely consequences of the unpredictability of bacterial activity in freshwater ecosystems after
treatment with allelochemicals and xenobiotics (EPA 2002;
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freshwater systems, the following information should be
determined: 1) the AA of xenobiotics and other chemicals
against the target alga; 2) the effects of these chemicals on
other organisms in the freshwater ecosystem; and 3) algal
dynamics after removal of the target alga. Most relevant studies conducted in Korea (Kim et al. 2003; Kim et al. 2006; Park
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rather than in open water. Technical limitations associated
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the water and sediment to sustain the cyanobacterial bloom.
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cyanobacterial blooms and to the determination of criteria for
the use of xenobiotics and other chemicals in research in situ.
The use of xenobiotics is often necessary, particularly in
preservation areas where there is a need to expedite water
treatment for biological remediation of algal bloom–affected
ecosystems.
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Submitted 3 March 2008
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Accepted 2 September 2008
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