Toxic Microcystis Aeruginosa Strains Under Oxidative Stress Conditions

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Comparative Growth Study of Toxic and Non-Toxic Microcystis aeruginosa Strains under Oxidative Stress Conditions

Neil Rajput

A PROJECT

submitted to

Oregon State University

University Honors College

in partial fulfillment of the requirement for the degree of

Honors Baccalaureate of Science in Microbiology (Honors Scholar)

Presented May 15, 2014 Commencement June 2014

AN ABSTRACT OF THE THESIS OF

Neil Rajput for the degree Honors Bachelor of Science in Microbiology presented on May 15, 2014. Title: Comparative Growth Study of Toxic and Non-Toxic Microcystis aeruginosa Strains under Oxidative Stress Conditions

Abstract approved: ______Dr. Theo Dreher, Mentor

Toxic cyanobacterial blooms in freshwater sources are of increasing concern due to the production of toxins that pose a threat to human health. Both toxic and non-toxic strains of Microcystis aeruginosa cohabitate with one-another, in vivo. However, environmental conditions play a large role in determining the dominance of toxic or non-toxic strains in a given cyanobacterial bloom. The mechanism underlying

Microcystis aeruginosa’s ability to out compete other strains in response to changing environmental conditions remains under investigation. This research studies the growth of different toxic and non-toxic strains of Microcystis aeruginosa under varying light intensities. Light intensity was analyzed as it was shown to be a source of photooxidative stress. A scopoletin assay was adapted to measure the evolution of hydrogen peroxide, which was found to be greater in samples exposed to high light treatment. The study also incorporated comparative genomics to highlight several conserved peroxiredoxin genes in certain strains of Microcystis aeruginosa that have been studied in other cyanobacterial species as stress-response mechanisms. This information was used to analyze and interpret environmental data published on ecological shifts of toxic potential that have been noted in many

Microcystis aeruginosa dominated blooms.

Key Words: Microcystis aeruginosa, oxidative stress, cyanobacterial blooms, scopoletin assay, microcystin, photooxidative stress, toxic strain dominance

Corresponding e-mail address: [email protected]

Honors Baccalaureate of Science in Microbiology project of Neil Rajput presented May 15, 2014.

APPROVED:

______Mentor, representing Microbiology

______Committee member, representing Microbiology

______Committee member, representing Biology

______Chair, Department of Microbiology

______Dean, University Honors College

I understand that my project will become a part of the permanent collection of Oregon State University, University Honors College. My signature below authorizes release of my project to any reader upon request.

Neil Rajput

ACKNOWLEDGEMENTS

I received support from many people over the course of my research work, which allowed me to write this thesis. First, I would like to thank the members of the Dr. Theo Dreher laboratory at Oregon State University for teaching me the “ins and outs” of laboratory research and for making me more familiar with techniques related to studying cyanobacterial species. They supported me in troubleshooting when parts of my experiment did not go as planned and their affable personalities made every day in the lab an exciting experience. I would like to thank Dr. Dreher for allowing me to do research in his lab and for his continual support and guidance in planning my research. A special thank you goes to my postgraduate doctoral mentor, Dr. Tim Otten, for helping me throughout the project and giving me insight through all stages of the research. Lastly, I would like to thank my advisors within both the University Honors College, and the Microbiology major for helping me plan my research and for overseeing my completion of the requirements of this thesis.

Thank you to my mom, dad, and sister for always supporting me throughout my undergraduate experience. Without their help, I wouldn’t be the person I am today. I thank them for inspiring me to pursue greater endeavors in life. I know that my next step in life, attending medical school, would not be possible without their support.

Funding for this project was provided by the Dr. Theo Dreher laboratory and all laboratory research related to this thesis was conducted in Corvallis, Oregon at

Oregon State University.

TABLE OF CONTENTS

INTRODUCTION 1 Background 1 Taxonomy 2 Microcystis aeruginosa: An Overview 3 Production of Photooxidative Stress 7 Population Dynamics 9

THESIS STATEMENT 10

MATERIALS AND METHODS 11 Determination of Light Conditions (Independent Variable) 11 Selection of Strains 12 Establishment of Log (Exponential Phase) Growth 12 Growing Conditions 13 Measurement of Photo-induced Oxidation (Scopoletin Assay) 13 Cell Counting Procedure 15 Comparative Genomics/Proteomics to Identify Presence and Function 16 of Peroxiredoxin Genes Evaluation and Comparison of Environmental Studies on the Variability 17 of Toxic Potential of Microcystis aeruginosa

RESULTS AND DISCUSSION 18 Day 1-14 Growth of Toxin and Non-Toxic Strains under High and Low 18 Light H2O2 Concentration Determination using Adapted Scopoletin Assay 22 Investigation of 2-cys Peroxiredoxin 27 Literature Review of Population Dynamics in Environmental Studies 29 (Bloom Shift from Toxic to Non-Toxic Strain Dominance) Misson, Benjamin, and Delphine Latour. 30 Zhu, Lin et al. 32 Van Wichelen, Jeroen et al. 33 Gobler, C. J. et al. 34 Yoshida, Mitsuhiro et al. 35 Further Support for Cyanophage-Mediated Genotypic Succession 36 of Microcystis aeruginosa over a Bloom Period

CONCLUSION 38

REFERENCES 40

LIST OF FIGURES

Figure

Figure 1. Structure of Microcystin-LR. Microcystin (mcy) gene cluster 4 Figure 2. Interaction of microcystin with phosphoprotein phosphatase (PPP) 6 protein that shows covalent binding to cysteine residues at various grooves. Figure 3. Enlarged Grid of Hemocytometer 16 Figure 4. Growth Curve (High Light / Low Light) Days 1-14. PCC 7005 - "Non- 20 Toxic" Figure 5. Growth Curve (High Light / Low Light) Days 1-14. PCC 2667 - "Toxic" 20 Figure 6. Growth Curve (High Light / Low Light) Days 1-14. CPCC 299 - "Toxic" 21

Figure 7. Growth Curve (High Light / Low Light) Days 1-14. UTEX 2386 - "Non- 21 Toxic"

Figure 8. Scopoletin Assay Standard Curve - Growth Day 11 24 Figure 9. Scopoletin Assay Standard Curve - Growth Day 14 24 Figure 10. BlastP Search of BAS1 (2-cys Prx) in Complete and Partially 28 Complete Genomes of Microcystis Aeruginosa Strains Figure 11. A Concept Map Depicting the Various Publications used in this 30 Literature Review and their Reported Mechanism for the Observed Genotypic Shift (Toxic to Non-Toxic) over a Bloom Period.

LIST OF TABLES

Table

Table 1. Scopoletin Assay for Determination of H2O2 Concentration - Day 11 22

Table 2. Scopoletin Assay for Determination of H2O2 Concentration - Day 14 23

Table 3. Difference in H2O2 Concentration between High Light and Low Light 25 (nM) - Growth Day 11

Table 4. Difference in H2O2 Concentration between High Light and Low Light 25 (nM) - Growth Day 14 Table 5. Amino Acid Sequence of 2-cys Peroxiredoxin (BAS1) 27

LIST OF EQUATIONS

Equation

Equation 1. Cell Density Equation for Hemocytometer Counting 15

DEDICATION

This thesis is dedicated to the professors and faculty

of the Microbiology department at Oregon State University

for their expertise and continual support.

Comparative Growth Study of Toxic and Non-Toxic Microcystis aeruginosa Strains

INTRODUCTION

Background

Toxic cyanobacterial blooms are of growing concern due to the environmental implications they have on water potability and overall watershed health. This issue is further complicated by rising global temperatures and alteration of seasonal weather patterns, which modify the temperature of water bodies leading to stratification (1). Studies investigating this phenomenon, in conjunction with eutrophication from freshwater pollution, have concluded that these changes will increase the level and severity of toxic cyanobacterial blooms globally (1).

Bloom formation is influenced by increasing global temperatures, agricultural run-off, and other sources that deliver abundant nitrogen and phosphorous that can be readily used by cyanobacterial species (2). Bloom formation poses significant threats to human and mammal health and to the health of other aquatic species. In humans, consumption of or swimming in water that is tainted with cyanobacteria is linked to gastroenteritis, skin irritation, and allergic reactions of the skin and eyes. Long-term exposure has been linked to liver damage

(3). The cause of these health concerns is due to cyanobacterial production of hepatotoxins, which interfere with eukaryotic cellular signaling (3). Cyanobacterial blooms also create issues for other aquatic life. When large areas of blooms die, the resulting organic matter favors the growth of other microbes. These microbes 2 deplete the waters of dissolved oxygen through their metabolic activity leading to hypoxic “dead-zones” where aquatic life cannot exist (4).

Taxonomy

The appearance of cyanobacterial species varies depending on species. They can exist unicellularly or alternatively in colonies. In colony form, they can form hollow balls or filaments, or sheets. Their appearance under a microscope is dominated by large gas vesicles, which confer buoyancy within the water column to many species (1). This allows them to take advantage of optimal light intensities and carbon dioxide levels by simply changing the volume of their gas vesicles; a process known as phototaxis (1). In comparison to other bacterial species, cyanobacteria are quite large, with sizes ranging from 1 to 10 μm.

Cyanobacteria are Gram-negative, photoautothophs, meaning they use light energy to drive metabolic processes and are capable of fixing carbon from an inorganic form (5). Their evolution as organisms is believed to have dramatically changed the composition of the earth’s environment by producing oxygen gas as a byproduct of photosynthesis, therefore stimulating a shift from a reducing environment to an oxidizing one (5). Research has indicated that cyanobacteria are the origin of chloroplasts found in plants and other eukaryotes, as described by the endosymbiotic theory (6).

Cyanobacterial photosynthesis is therefore somewhat similar to that of plants. Cyanobacteria use a pigment known as phycocyaninin in addition to 3 chlorophyll-a to capture light, which gives them their blue-green appearance (7).

Water is used as an electron donor and oxygen is produced as a product.

Cyanobacteria perform photosynthesis via photosystem (PS) II and I, which allows water to be oxidized through Z-scheme (7). Phycocyanin and chlorophyll-a are housed in thylakoid membranes. PS II captures light energy, which initiates an electron transport chain. The Calvin Cycle process allows carbon dioxide to be fixed into organic carbon (7).

Microcystis aeruginosa: An overview

Microcystis aeruginosa is a member of the phylum, Cyanobacteria, that is found in freshwater sources. Microcystis aeruginosa is one of the most common cyanobacterial species related to eutrophic blooms (4). Microcystis aeruginosa exists in vivo in both toxic and non-toxic strains. In toxic strains, Microcystis aeruginosa produces microcystin, a cyclic peptide hepatotoxin that is harmful to human and mammalian health. Microcystin is produced as a secondary metabolite, and is not directly related to Microcystis aeruginosa’s key biochemical pathways leading to growth or survival (8). Microcystis aeruginosa can also produce lipopolysaccharide, a skin irritant (14).

Over 80 variants of the microcystin toxin have been isolated from cyanobacterial species (13). Microcystins differ from each other by the differentiation of L-amino acids located in two variable sites, and sometimes by the chemical modification of other amino acids in the protein. Each microcystin has a different toxicity profile; however, microcystin-LR is the most toxic and most 4 studied variety of microcystins. In Microcystis aeruginosa, microcystin-LR is encoded by a 55 kb gene cluster, mcy. Mcy is subdivided into 6 gene sections that encode larger proteins mcyA-E and mcyG, which have polyketide synthase activity and nonribosomal peptide synthetase activity (13). mcy also encodes smaller proteins mcyF and mcyH-J. All microcystins are non-ribosomal peptides meaning that they are not synthesized by ribosomes (13). The different protein “modules” on the larger proteins provide specialized enzymatic function that allows for biosynthesis of the peptide.

Figure 1. Structure of Microcystin-LR. (above). Microcystin (mcy) gene cluster (below). (13). 5

Eutrophication results from agricultural run-off and other sources, resulting in excess nitrogen and phosphorus (1, 14, 21). These nutrients, when in excess, allow Microcystis aeruginosa to greatly proliferate. Microcystis aeruginosa also undergoes high levels of photosynthesis. This allows Microcystis aeruginosa to dominate cyanobacterial blooms under typical bloom forming conditions (1). Heavy bloom formation typically begins in the early summer and may last 2-4 months.

Microcystin exposure is a concern because typical processes to treat water for potability do not effectively remove microcystin (3).

Microcystin’s toxicity affects hepatocytes (liver cells), which decreases the liver’s function. Microcystin-LR’s interaction with the phosphatases causes covalent bond formation between the methylene group of microcystin and a cysteine residue on the phosphoprotein phosphatase (PPP) (46) (Figure 2). Microcystin covalently binds to the hydrophobic groove, acidic groove, and C-terminal groove in a “Y” configuration. This bond blocks access of the substrate to the active site of the PPP enzyme, rendering PPP nonfunctional and preventing phosphorylated proteins from becoming de-phosphorylated (46). More specifically, the toxin works as an inhibitor of mammalian phosphatase 1 and 2A (types of PPP), and an activator of cycloxygenase and phospholipase A2. This causes hyper-phosphorylation of cytokeratin proteins that leads to changes in cell shape and rearrangement of intermediate filaments of liver cells. The destruction of liver cells leads to hepatic hemorrhage or hepatic insufficiency. 6

More recently, microcystin was found to interact with the mitochondria of mammalian cells. Here, microcystin is believed to disrupt normal calcium (Ca2+) signaling in the mitochondrial membrane leading to membrane instability and induction of reactive oxygen species (ROS). The accumulation of high concentrations of ROS leads to apoptosis of the mammalian cell. This is a more recently proposed mechanism for microcystin’s toxicity and it is still under investigation (45).

Figure 2. Interaction of microcystin with phosphoprotein phosphatase (PPP) protein that shows covalent binding to cysteine residues at various grooves. (46)

Microcystin role in Microcystis aeruginosa’s fitness has not been elucidated.

The evolutionary advantage for Microcystis aeruginosa is unclear because the organism only releases significant amounts of microcystin following cell death and lysis (8). A study done by Rohrlack et al. on the survival of Daphnia (a grazing zooplankton) failed to link the toxin as the cause of decline in feeding Daphnia 7 populations (29). Furthermore, evolutionary history has revealed that mcy genes that synthesize microcystin evolved with other housekeeping genes well before the appearance of eukaryotic grazers (such as metazoans) (28), therefore reducing support for the theory that microcystin production is a defense mechanism to inhibit zooplankton grazing. Another study conducted by Schatz provided insight that microcystin production might be used as intercellular signaling, inducing apoptosis to send signals to the surrounding population warning of stress (30). A study done by Börner found that microcystin (mcy) genes underwent greater transcription into mRNA following exposure to high light, although the total quantity of microcystin did not increase (31). A proteomic analysis in a study by

Zilliges et al. revealed that microcystin may engage in binding to cysteine residues on enzymes of the Calvin cycle, phycobiliproteins and NADPH-dependent reductases, thereby stabilizing these proteins during high light and oxidative stress, indicating that microcystin may function in the tolerance of oxidative stress (8).

Prior findings that microcystin acted as a siderophore to sequester iron in low-iron conditions were dismissed by a recent study by Klein (15). This study found that similar strains of Microcystis aeruginosa have the same amount of iron uptake despite one of the strains having a gene knockout for mcy.

Production of Photooxidative Stress

High light intensity on organic matter can lead to the production of reactive oxygen species, which in an aquatic environment leads to an increase of hydrogen peroxide (H2O2) concentration (7). Hydrogen peroxide, a strong oxidizing agent is 8 particularly damaging to enzymes involved in photosynthesis and the Calvin Cycle, thereby leading to an increase in stress on aquatic organisms (43). Hydrogen peroxide has also been shown to disrupt the cellular membrane, as well as cause atypical DNA laddering leading to apoptosis (44). Absorption of high levels of solar radiation increases the rate of photosynthetic electron transport relative to the rate of electron consumption during CO2 fixation (32). The result of this imbalance is partially reduced forms of oxygen (reactive oxygen species), which are in between atmospheric oxygen (most oxidized form) and water (most reduced form), in terms of redox state, such as O2- (7). These forms of oxygen can react with water to create hydrogen peroxide, which is soluble in and can cross the cellular membrane.

Excessive reactive oxygen species in cyanobacteria can lead to photosystem II inactivation, protein and nucleic acid damage, and therefore leads to growth inhibition and death (43). A study by Ding et al. found that exposing Microcystis aeruginosa cells to 250 and 325 µM H2O2 showed membrane deformation and partial disintegration of thylakoids (visualized with electron microscopy).

Photosynthetic efficiency, measured through the ratio of variable fluorescence to maximum fluorescence, and the maximum electron transport rate, were also significantly decreased after exposure to high concentrations of hydrogen peroxide

(44). Zilleges et al. noted that microcystin may function in binding to RbcL, RbcS and

Prk gene products, which are subunits of the Calvin Cycle protein, RubisCO. In binding to RubisCO, microcystin may help stabilize it from becoming oxidized/denatured during high oxidative (high light) conditions (8).

Population Dynamics

In cyanobacterial bloom populations, both toxic and non-toxic Microcystis aeruginosa strains can exist with one another, but studies on population dynamics have reported shifts in the dominance of toxic and non-toxic strains over a bloom season (11, 21, 22, 23, 40, 41, 42). Research investigating bloom population dynamics and its causes is still an active area of research among aquatic microbial ecology.

THESIS STATEMENT

Exposing Microcystis aeruginosa cultures to high light conditions induces greater oxidative stress to the organism, which may be detected using an adapted scopoletin-based assay. The hypothesis guiding this study is that this photooxidative environmental stress is combated by the presence of microcystin and peroxiredoxins, providing insight into the dominance of toxic strains over non-toxic strains in vivo.

MATERIALS AND METHODS

Determination of Light Conditions (Independent Variable)

To assess what intensities of light would be effective to use as high light and low light (control) conditions, environmental data and prior studies were consulted. Microcystis aeruginosa strain PCC 7806 has been found to grow optimally in light with photosynthetically active radiation (PAR) value at 40 μmol of photons m−2 s−1. PAR values above 80 μmol of photons m−2 s−1 showed a decrease in the strain's normal growth rate (9). Another study used values of 20 μmol of photons m−2 s−1 to represent control light conditions, and noted significant stress on growing cultures at a PAR value of 300 μmol of photons m−2 s−1 (1). For this study, a PAR of

30 μmol of photons m−2 s−1 was selected to use as low light conditions, and 300 μmol of photons m−2 s−1 was selected to use as high light treatment. The rationale was to have the low light treatment reflect normal Microcystis aeruginosa growth, which would represent a scientific control. The high light PAR was chosen to induce significant stress on the actively growing cultures and to use enough intensity of light for the assessment of the generation of photooxidative stress produced by light on organic matter. To achieve these values, a light bank was set up using fluorescent bulbs. Using a light meter, the distance between the light bulbs and the cultures to be grown in flasks was adjusted to achieve desired light intensities. The tops of the flasks for the low light treatment were placed approximately 40 centimeters from the tops of the light bank and tops of the flasks for the high light treatment were placed 10 centimeters from the light bank. These distances provided the PARs 12 desired for use in this experiment. The light banks were set on a timer to achieve 12 hours of light and 12 hours of darkness, to simulate natural conditions and to allow the cultures to carry out a normal cycle of photosynthesis and cellular respiration per day.

Selection of Strains

Two toxic strains and two non-toxic strains of Microcystis aeruginosa were selected for this study. The strains used were: UTEX 2667 and CPCC 299

(toxic) and PCC 7005 and UTEX 2386 (non-toxic). These strains were obtained from stock cultures maintained in the Dreher laboratory. Strains were selected based on their availability to use in the experiment, and to also represent strains of

Microcystis aeruginosa from different geographical locations. UTEX 2667 and UTEX

2386 are culture strain from a collection at University of Texas at Austin originally isolated from Little Rideau Lake, Ontario, Canada. PCC 7005 was obtained from the

Pasteur Culture Collection.

Establishment of Log (Exponential Phase) Growth

In order to use actively growing cells in exponential phase for this experiment, samples of UTEX 2667, CPCC 299, PCC 7005, and UTEX 2386 were taken from stock cultures and transferred to sterile 75-ml Corning plastic flasks containing BG11 medium (10). These cells were grown at standard conditions of 23°

Celsius and a PAR of 30 μmol of photons m−2 s−1. Cell density was assessed at 1, 3, 5 and 7-day time points and plotted to visualize the growth curve of the strains. This 13 was done using a hemocytometer counting procedure for Microcystis aeruginosa. On day 7, the cells of all four strains were determined to be in log (exponential growth) and were deemed suitable to transfer and use for further experimentation. Log

(exponential growth) was assessed through a visual assessment of the curve made from plotting cellular density over time.

Growing Conditions

For the low light and high light treatments on UTEX 2667, CPCC 299, PCC

7005, and UTEX 2386, the initial cell density for Day 1 starting values was established by transferring cells from log phase growth to new BG11 medium in 75- mL Corning plastic flasks. This was done by establishing the cellular density of the transferrant through hemocytometer counting. Once cellular density values of the log phase cells were determined, a volume of cell suspension was removed via pipette. Culture samples of 60 ml with a cellular density of 4.0 x 106 cells/ml were achieved though dilution of the log phase cells. This was the starting density used in the low and high light experiment. It was important that cell cultures used in the experiment began at the same cellular density so that changes in growth over the course of the experiment could be more readily compared.

Measurement of Photo-induced Oxidation (Scopoletin Assay)

To measure the amount of photooxidation that occurred as a result of the high light treatment, a special procedure involving scopoletin had to be adapted.

The scopoletin procedure was used because no formal scientific instruments could 14 be obtained to measure hydrogen peroxide concentration in solution. Furthermore, there is no known procedure that has been described using scopoletin to study hydrogen peroxide concentration in samples of cyanobacteria. As a result, the procedure used in this experiment was an adaptation of a procedures published by

Kieber and Corbett (33, 17). Kieber et al. studied hydrogen peroxide production in natural waters from samples collected from Paint Branch, a stream in Maryland

(33). Scopoletin can be used to measure hydrogen peroxide because scopoletin is a naturally fluorescent substrate. Horseradish peroxidase catalyzes the oxidation of scopoletin by hydrogen peroxide, therefore decreasing scopoletin’s fluorescence

(33). A borate buffer stops the reaction so that the fluorescence can be measured by a fluorometer at a wavelength that is able to detect scopoletin fluorescence (33).

Therefore scopoletin’s measured fluorescence in the assay is inversely proportional to the amount of hydrogen peroxide present in the sample (17). In this experiment,

60 μL of sample was added to 20 μL of 0.2 M sodium acetate / 1 nM EDTA, pH 4.7.

10 μL of horseradish peroxidase (30 μg/ml) was added, followed by 10 μL scopoletin (0.04 mM, 770 ng/ml). The reaction was incubated for 10 minutes, and

100 μL of 0.15 M potassium borate, pH 10, was used to stop the reaction. The assay was performed in a 96 well black Corning plate with a clear bottom. The fluorescence was read in a Tecan Infinite 200 plate reader (with fluorometer function) at an excitation wavelength of 395 nm, and an emission wavelength of 470 nm, at a gain of 150. A standard curve of scopoletin fluorescence was generated using known concentrations of hydrogen peroxide that were prepared via serial 15

dilutions (2.0 μM, 1.0 μM, 500 nM, 250 nM, 100 nM, 50 nM H2O2). This curve provided a reference for experimental values to be deduced.

Cell Counting Procedure

Cell density at various time points during the trials was measured by using a light microscope and a hemocytometer slide. This is a common method for assessing cyanobacterial concentrations in samples. Because of Microcystis aeruginosa’s deep pigment, no staining was needed to view the cells under the microscope. Cells were counted in a grid in which each square represents a certain volume, in this case 0.1 mm3. For statistical accuracy, if cells were positioned on the gridlines of the counting field, they were only counted on two of the four sides. An equation

(Equation 1) could be used to calculate cell density (cells/ml) based on the average number of cells counted in a 0.1 mm3 volume.

Equation 1. Cell Density Equation for Hemocytometer Counting

average cell count per 1mm square * 2 * 104 = Viable cell count (cells/ml)

If the number of counts exceeded 200 then a dilution of the sample was done and this was considered in the calculations. If the number of counts were less than 50, then a greater area of the hemocytometer was counted and this was also reflected in the calculations. Cells were not counted on two of the four edges for statistical accuracy (Figure 3). Furthermore, for better statistical accuracy, 5 grids on the hemocytometer were counted and these values were averaged. Replicate counts using the same procedure were also obtained for each assessment of cellular density. 16

Figure 3. Enlarged Grid of Hemocytometer (note: counting procedure)

Comparative Genomics/Proteomics to Identify Presence and Function of Peroxiredoxin Genes

Despite several studies done on proposed mechanisms on how Microcystis aeruginosa deals with oxidative stress, none of these studies have investigated if peroxiredoxins are involved in Microcystis aeruginosa’s stress response. 2-cys peroxiredoxin has been studied as a possible protein that functions in the oxidative stress response in Listeria monocytogenes. (34) The prevalence of peroxiredoxin genes was assessed in different strains of Microcystis aeruginosa using a genomics search engine and a literature review of peroxiredoxin function was done to better hypothesize the protein’s role in managing oxidative stress.

Evaluation and Comparison of Environmental Studies on the Variability of Toxic Potential of Microcystis aeruginosa

Several environmental studies focused on characterizing population dynamics in cyanobacterial blooms have noted that many blooms experience genotypic structure shifts in population from being heavily toxic in the beginning

(having a high percentage of microcystin producing strains) to a higher dominance of non-toxin strains after the bloom’s peak. Several mechanisms for this have been proposed. A literature review was done to compare various findings among different environmental studies.

RESULTS AND DISCUSSION

Day 1-14 Growth of Toxin and Non-Toxic Strains under High and Low Light

The 14 day growth studies on UTEX 2667 and CPCC 299 (toxic) and PCC

7005 and UTEX 2386 (non-toxic) strains all began at the same initial concentration of 4.0 x 106 cells/ml which were transferred from stock cultures containing cells in log phase. In all four of the strains examined, the low light conditions produced a greater amount of growth (assessed though Day 14 end point growth values) than the high light conditions (Figures 4-7). This result was consistent with the hypothesis that high light would induce a greater level of stress of the samples. The high light growth at day 14 of UTEX 2667 (toxic) was 75.53% of the low light growth value (Figure 5). The high light growth at day 14 of CPCC 299 (toxic) was

86.72% of the low light growth value (Figure 6). For the non-toxic strains, the percentage of high light growth compared to low light growth was significantly less:

61.82% for PCC 7005, and 56.20% for UTEX 2386, respectively (Figures 4, 7). In is noted in Zilliges et al. that microcystin may function in binding to RbcL, RbcS and

Prk gene products that are subunits of the Calvin Cycle protein, RubisCO. In binding to RubisCO, microcystin may help stabilize it from becoming oxidized/denatured during high oxidative (high light) conditions (8). This mechanism could potentially provide insight into why the toxic strains investigated achieved higher high light to low light growth ratios than that of the non-toxic strains investigated. Lastly, the difference in growth rate between strains is likely attributable to natural features among strains (i.e. some strains may grow faster than others). These features were not the subject of investigation in this study. 19

It is important to note that innate genotypic variations between strains may cause some strains to have faster doubling times and a higher rate of growth than other strains. One concern about the results is that the growth of strains UTEX 2667

(Figure 5) and CPCC 299 (Figure 6) is overall less than strains PCC 7005 (Figure 4) and UTEX 2386 (Figure 7). It may be that each strain has specific media requirements or optimal control growth PAR that was different than those used in this study. In this study BG11 media was used which is a “complete” media for growing cyanobacterial species, meaning that it is not deficient in any nutrients for

Microcystis aeruginosa’s normal metabolism. The lighting conditions used in this study could also have influenced the growth of the strains and could be a possible reason for the lower growth rate of UTEX 2667 (Figure 5) and CPCC 299 (Figure 6).

As noted, some strains may achieve their optimal growth rate at a different intensity of light than other strains. In this study a PAR value of 30 μmol of photons m−2 s−1 was used as the control light condition; however it may be possible that strains

UTEX 2667 and CPCC 299 grow better under different PAR intensities. Perhaps a different and individualized PAR “control” value is needed for each strain, which could allow for a more valid comparison of the results (from strain to strain).

Further testing on the strain’s individualized growth patterns under different light intensities would provide further insight to clarify this potential shortcoming.

Microcystis aeruginosa Strain PCC 7005 - "Non-Toxic" 4.00E+07 3.50E+07 3.00E+07 2.50E+07 2.00E+07 1.50E+07 1.00E+07 5.00E+06 Cell Density (cells/ml) 0.00E+00 0 2 4 6 8 10 12 14 16 Day

LL HL

Figure 4. Growth Curve (High Light / Low Light) Days 1-14. PCC 7005 - "Non-Toxic"

Microcystis aeruginosa Strain UTEX 2667 - "Toxic" 4.00E+07 3.50E+07 3.00E+07 2.50E+07 2.00E+07 1.50E+07 1.00E+07 5.00E+06 Cell Density (cells/ml) 0.00E+00 0 2 4 6 8 10 12 14 16 Day

LL HL

Figure 5. Growth Curve (High Light / Low Light) Days 1-14. PCC 2667 - "Toxic"

Microcystis aeruginosa Strain CPCC 299 - "Toxic" 4.00E+07 3.50E+07 3.00E+07 2.50E+07 2.00E+07 1.50E+07 1.00E+07 5.00E+06 Cell Density (cells/ml) 0.00E+00 0 2 4 6 8 10 12 14 16 Day

LL HL

Figure 6. Growth Curve (High Light / Low Light) Days 1-14. CPCC 299 - "Toxic"

Microcystis aeruginosa Strain UTEX 2386 - "Non-Toxic" 4.00E+07 3.50E+07 3.00E+07 2.50E+07 2.00E+07 1.50E+07 1.00E+07 5.00E+06 Cell Density (cells/ml) 0.00E+00 0 2 4 6 8 10 12 14 16 Day

LL HL

Figure 7. Growth Curve (High Light / Low Light) Days 1-14. UTEX 2386 - "Non-Toxic"

H2O2 Concentration Determination using Adapted Scopoletin Assay

Table 1. Scopoletin Assay for Determination of H2O2 Concentration - Day 11*

Experimentally Derived Sample Fluorescence Concentrations of H2O2 (nM) Catalase- treated 20504 Water

(Control)

BG-11 19932 (Media)

2.0 μM H2O2 4309 1.0 μM H2O2 12645 500 nM H2O2 16102

250 nM H2O2 18584

100 nM H2O2 19120

50 nM H2O2 20073 PCC 7005 HL 18960 174.30 PCC 7005 18103 281.69 HL/S PCC 7005 LL 19168 148.24 PCC 7005 18240 264.53 LL/S UTEX 2667 19108 155.76 HL UTEX 2667 19309 130.57 LL CPCC 299 HL 19199 144.36 CPCC 299 LL 19403 118.79 UTEX 2386 19202 143.98 HL UTEX 2386 19337 127.06 LL *average background fluorescence subtracted

Table 2. Scopoletin Assay for Determination of H2O2 Concentration - Day 14*

Experimentally Derived Sample Fluorescence Concentrations of H2O2 (nM) Catalase- treated 20104 Water

(Control)

BG-11 19787 (Media)

2.0 μM H2O2 4109 1.0 μM H2O2 12365 500 nM H2O2 16350

250 nM H2O2 18102

100 nM H2O2 18887

50 nM H2O2 19666 PCC 7005 HL 19201 108.12 PCC 7005 18374 212.82 HL/S PCC 7005 LL 19377 85.84 PCC 7005 18689 172.94 LL/S UTEX 2667 19200 108.25 HL UTEX 2667 19433 78.75 LL CPCC 299 HL 18855 151.92 CPCC 299 LL 19116 118.88 UTEX 2386 18795 159.52 HL UTEX 2386 19082 123.18 LL *average background fluorescence subtracted

Scopoletin Assay Standard Curve - Growth Day 11 25000

20000

15000

10000

Units Fluorescence 5000 y = -7.8987x + 20055

0 0 500 1000 1500 2000 2500 H2O2 Concentration (nM)

Figure 8. Scopoletin Assay Standard Curve - Growth Day 11

Scopoletin Assay Standard Curve - Growth Day 14 25000

20000

15000

10000

Units Fluorescence 5000 y = -7.9803x + 20351

0 0 500 1000 1500 2000 2500 H2O2 Concentration (nM)

Figure 9. Scopoletin Assay Standard Curve - Growth Day 14

Table 3. Difference in H2O2 Concentration between High Light and Low Light (nM) - Growth Day 11

Difference in H2O2 Strain Concentration between HL and LL (nM) PCC 7005 (MC-) 26.06 UTEX 2667 (MC+) 25.19 CPCC 299 (MC+) 25.56 UTEX 2386 (MC-) 16.92

Table 4. Difference in H2O2 Concentration between High Light and Low Light (nM) - Growth Day 14

Difference in H2O2 Strain Concentration between HL and LL (nM) PCC 7005 (MC-) 22.28 UTEX 2667 (MC+) 29.50 CPCC 299 (MC-) 33.04 UTEX 2386 (MC+) 36.34

The scopoletin assay allowed for experimental H2O2 concentrations to be derived. Fluorescence values for the samples were read in a fluorometer, which quantified the amount of fluorescence in the sample. A standard curve was generated so that experimental values could be derived (Figures 8 and 9). When plotted, the standard curve gave values that followed a linear pattern. Using the equation of this line, the experimental concentrations were determined by calculation. The values listed in Tables 1 and 2 represent the average of two replicates. The average background fluorescence of the plate was subtracted to ensure that values represented only the fluorescence of the scopoletin in the assay.

The results of the scopoletin assay indicate that on Day 11 toxic and non- toxic high light samples had a greater H2O2 concentration than low light samples 26

(between 16.92 and 26.06 nM higher) (Table 3). A similar result was obtained by

Day 14 samples; the high light samples showed a greater H2O2 concentration than low light samples (between 22.28 and 36.34 nM higher) (Table 4). These results were consistent with the hypothesis that exposing the cell cultures to high light would induce the generation of more reactive oxygen species than exposing them to low light (due to the overstimulation of the electron transport chain machinery during photosynthesis, which creates reactive oxygen species) (7). These results are also consistent with findings by Cooper et al. in a study that exposed natural waters to high light and observed similarly increased concentrations of H2O2 (20). The experimentally derived values are also similar to those investigated in a study by

Leunert et al. (16). This study had hypothesized that Day 14 derived H2O2 concentrations would be greater than Day 11 derived concentrations (due to the increase in organic matter caused by an increase in cellular density). However, this particular hypothesis is not confirmed by the results, as several of the strain’s Day

11 H2O2 concentrations are higher than their respective values at Day 14.

Investigation of 2-cys Peroxiredoxin

Through background research on the effects of oxidative stress on cellular function, it was expected that peroxiredoxins play a role in an organism’s ability to control intracellular peroxide levels. Peroxiredoxins function by having a conserved

“peroxidatic cysteine”(CP) residue which readily reduces peroxide substrates. The mechanism is as follows: when a peroxidatic cysteine is oxidized by a peroxide, a conformation change occurs around the active site of the enzyme, which allows a free thiol to form a disulfide bond with the CP (27). The class 2-cys prx refers to a peroxiredoxin in which the thiol group is on the peroxiredoxin (27).

Despite extensive literary research, there was little to no published work of the role of peroxiredoxins specific to Microcystis aeruginosa. A putative sequence of 2- cys-prx was annotated in the complete genome sequence of Microcystis aeruginosa

TAIHU98, a nontoxic strain isolated from Taihu Lake in China (Table 5) (19). A

BlastP search revealed this 2-cys peroxiredoxin “BAS1” was present in 14 other partially or wholly sequence strains of Microcystis aeruginosa (Figure 10).

Table 5. Amino Acid Sequence of 2-cys Peroxiredoxin (BAS1) (19)

>tr|L7E2N1|L7E2N1_MICAE 2-Cys peroxiredoxin BAS1 OS=Microcystis aeruginosa TAIHU98 GN=O53_2506 PE=4 SV=1

MTAEGCLRVGQAAPDFTATAVFDQEFKTIKLSDYRGKYVVLFFYPLDFTFVCPTEITAFS DRVSEFASINTEILGVSVDSEFAHLAWIQTERKSGGVGDVAYPLVSDLKKEISTAYNVLD PDAGVSLRGLFIIDKEGVIQHATINNLSFGRSVDETLRTLKAIQYVQSHPDEVCPAGWQE GDATMVPDPVKSKVYFAAV

Figure 10. BlastP Search of BAS1 (2-cys Prx) in Complete and Partially Complete Genomes of Microcystis aeruginosa Strains

A study on assessing the presence of putative peroxiredoxins among the phylum,

Cyanobacteria, found that Microcystis aeruginosa strain NIES-843 contains seven putative prx genes (35). The functions of these genes and their expression profile have not been fully investigated. Two separate studies on Synechocystis sp. PCC

6803 and Synechococcus sp. PCC 7942 found that the disruption of genes encoding

2-cys Prx affected the strains’ tolerance to oxidative stress (25, 26). A more recent study looked at differences in the transcriptome between night-time and day-time periods in a strain of Microcystis aeruginosa derived from an environmental sample in Singapore (16). The results of this study indicated that a gene for peroxiredoxin was transcribed at a higher rate during daytime hours than at night-time hours. This suggests that the transcription of peroxiredoxin genes may be related to the presence of light or when the cell is actively photosynthesizing. Overall, it appears that the function of 2-cys peroxiredoxin in Microcystis aeruginosa is a topic in need of further investigation. 29

Literature Review of Population Dynamics in Environmental Studies (Bloom Shift from Toxic to Non-Toxic Strain Dominance)

Despite knowledge of biotic and abiotic factors that affect Microcystis aeruginosa’s growth (nutrient availability, temperature, salinity, etc.), the causes for temporal changes in genotypes have yet to be elucidated. A 2007 study by Kardinaal et al. was one of the first publications to report on a genotypic shift from toxic to non-toxic dominance over the course of a bloom. This study investigated hypertrophic lakes in the western region of The Netherlands (22). Populations were monitored using the rRNA of the internal transcribed spacer (ITS) region in combination with denaturing gradient gel electrophoresis (DGGE). For Microcystis aeruginosa, microcystin-producing and non-producing colonies were separated into different rRNA-ITS classes. The results of the study showed a seasonal succession of the Microcystis genotype in all three lakes, with a stronger presence of non-toxic genotypes after the peak of the bloom (towards the end of the season). Despite these findings, the study did not investigate a mechanism for the observed shift. A

2010 study by Bozarth et al. also observed Microcystis strain successions within a single bloom period with surface samples from the Copco Reservoir in Northern

California from a 2007 bloom. These samples were analyzed genetically by sequencing clone libraries made with amplicons of the internal transcribed spacer of the rRNA operon (ITS), cpcBA, and mcyA. The study reached similar findings as in the study by Kardinaal et al. and also noted that the cause of this shift is not fully understood (41). Based on these studies, it appears that a common pattern in

Microcystis aeruginosa blooms is a shift from toxic to non-toxic dominance mid- 30 season. Here we compare and contrast various published work on the topic (Figure

11).

Figure 11. A Concept Map Depicting the Various Publications used in this Literature Review and their Reported Mechanism for the Observed Genotypic Shift (Toxic to Non-Toxic) over a Bloom Period

Misson, Benjamin, and Delphine Latour. "Vertical Heterogeneity of Genotypic Structure and Toxic Potential within Populations of the Harmful Cyanobacterium Microcystis aeruginosa."

Mission et al. studied Microcystis aeruginosa-dominated blooms in lakes

Grangent and Villerest, two artificial dam reservoirs of the Loire River (France) (11).

The researchers collected samples from three blooms at various depths: −0.5m,

−2.5m, −5m, −7.5m, −10m, −15m, -20 m, −25m and −30m. Cell enumeration was done by counting under a light microscope. A quantitative qPCR assay was developed to amplify the mcyB gene, which encodes an essential protein needed for 31 microcystin biosynthesis. This method allowed for the estimation of the proportion of toxic cells of Microcystis aeruginosa in the sample. The results of the study found that blooms of Microcystis aeruginosa can differ in genotype and toxic-potential in the vertical (depth) dimension. This vertical genotypic variation occurred predominantly in shallower depths of water. Interestingly, the researchers found that vertical differences in genotype were sometimes greater than horizontal differences in genotype (site samples at various locations at the same depth). The researchers also found that over the course of a bloom season, genotypic shifts occur among strains occupying certain depths of the water column. The researchers believed a driving factor behind the heterogeneity and observed shifts is the light attenuation in the deeper parts of the water column. They concluded that certain non-toxic/toxic strains are better suited for growth at different light intensities than others, and that depending on environmental conditions, both toxic and non-toxic strains undergo light-induced taxis by regulating the volume of their gas vacuole.

The study also noted a potential concern for environmental research work on population dynamics in cyanobacterial blooms, primarily concerning sampling method. The study suggests that if samples are collected from inconsistent depths over the course of a bloom, the innate vertical genotypic variation within the water column may produce sample isolates with different strains, thus giving the false illusion that genotypic succession is occurring. The study alluded to the possibility that improper sampling technique could skew the findings of other studies that have reported observed genotypic succession. This is an important finding because it provides a previously unidentified aspect for assessing prior studies, and provides 32 further emphasis on the importance of sampling depth for future studies on the topic.

Zhu, Lin et al. "Ecological dynamics of toxic Microcystis spp. and microcystin- degrading bacteria in Dianchi Lake, China."

A study by Zhu et al. investigated Dianchi Lake, a freshwater lake located in

Yunnan Province, China (23). Samples were taken between June 2010 and

December 2011 at three sampling sites (monthly). Quantitative-PCR was used to measure changes in the population of toxic and non-toxic Microcystis aeruginosa over the bloom period. qPCR was also used to study the expression of mlrA, a gene in microcystin-degrading bacteria (class Alphaproteobacteria, order

Sphingomonadales) that encodes an enzyme responsible for the hydrolytic cleavage of the cyclic structure of microcystins. The results of the study indicated that peaks in the microcystin concentration were apparent in September 2010 and October

2011, which was followed by peaks in the mlrA gene copy numbers of MC-degrading bacteria. The peak in mlrA gene copy number appeared one month after both annual peaks in microcystin (October 2010 and November 2011). September 2010 concentrations of microcystin were 1.33 g/liter at site D13, 1.3 g/liter at site D22, and 1.63 g/liter at site D24; October 2011 concentrations were 1.421 g/liter at site

D13, 1.39 g/liter at site D22, and 1.53 g/liter at site D24). The study found the proportion of toxic Microcystis cells in the lake varied from 93.8% to 2.9%, reaching a maximum in June and July, and a low in September to April. The researchers found the largest Microcystis blooms always occurred from June to December (108 copies/liter to 109 copies/liter), while the smallest blooms occurred from February 33 to April (106 copies/liter). The non-toxic strain appeared to be dominant between

September and December. One possible issue with this study is the amount of time elapsed between samples which may not allow for the determination of actual

“peak” values as the study indicated. These results of this study are important for inclusion in this literature review because they provide the basis for further discussion. While it is noted that peaks in microcystin-degrading bacteria numbers followed peaks in microcystin quantity, the research fails to provide insight into the root cause of microcystin release. Microcystin release in the extracellular environment is typically highest when cell death and apoptosis occurs. This research fails to find any support that microcystin-degrading bacteria are associated with cell apoptosis or the observed genotypic shift from toxic to non-toxic strains during bloom periods. This research may be of more significance if the MC- degrading bacterium cited in the study (genus Sphingopyxis) was found to be a predatory bacterium. Predatory bacteria are bacteria which are able to infect and feed off of other live bacterial cells. However, further investigation into predatory and potentially predatory signatures (based on certain genomic features) could not confirm or deny that any species in genus Sphingopyxis are predatory in nature (47).

Van Wichelen, Jeroen et al. “Strong effects of amoebae grazing on the biomass and genetic structure of a Microcystis bloom (Cyanobacteria).”

In a research publication by Van Wichelen et al., a two-year study was done on a small hypertrophic pond (37). Microcystis populations were enumerated through microscopy and DGGE of the ITS rDNA region was used to assess population dynamics. ITS-DGGE allows for the differentiation of closely related 34 organisms. This high-resolution method allows for the monitoring of population dynamics, by giving unique DGGE profiles even in comparison to closely related organisms. The study revealed that amoebae grazing affected the population dynamics of Microcystis and resulted in significant bloom biomass reduction in both years of the study. Grazing experiments revealed that amoebae (Genus Protozoa) had a preference for Microcystis aeruginosa compared to Microcystis viridis. This was shown to drive a shift in one of the bloom years from Microcystis aeruginosa dominance to Microcystis viridis dominance. The importance of this study is the revelation that grazers may have a preference for whether or not the bloom is producing toxin. This may have implications in genotypic shifts in blooms.

Gobler, C. J. et al. "Interactive influences of nutrient loading, zooplankton grazing, and microcystin synthetase gene expression on cyanobacterial bloom dynamics in a eutrophic New York lake."

A study by Gobler et al. evaluated the effects of zooplankton

(mesozooplankton and microzooplankton) and the population dynamics and toxin production of a bloom in Lake Agawam, a eutrophic lake in New York (40). This bloom was mainly dominated by toxic Microcystis aeruginosa in its onset. The research team observed that nitrogen levels enhance the growth rate and toxin levels produced by a toxic-strain dominated bloom. The team also observed the mesoplankton grazing was unable to reduce the bloom size when cell numbers were above a threshold of 8.0 x 104 cells/mL. The team proposed that both microcystin synthase gene expression and high cell densities under nutrient loading are mechanisms to deter grazing by zooplankton. The team observed that by late 35 summer, and early fall there was a demise in the bloom, caused by nitrogen- limitation which causes bloom cells to go into stationary phase. The team observed that cells in this stage of growth have lower expression of the microcystin synthase gene, which leads them to be more vulnerable to grazing by mesozooplankton leading to a rapid decline in the bloom. This decline could provide a niche for non- toxic Microcystis aeruginosa to fill and dominate. Despite this, the research fails to address what mechanisms the non-toxic strains have to inhibit mesoplankton grazing.

The study by Gobler et al. differs with the findings of another study by Li et al. that investigated nitrogen concentration’s effects on population dynamics. This study used a quantitative (real-time) PCR assay of the phycocyanin intergenic spacer (PC-IGS) and mcyD to study population dynamics in 2009 and 2010 samples from Lake Taihu (China) (36). The findings of this study were that the abundance of toxic and potentially-toxic (strains with the microcystin gene cluster in their genome) had a negative correlation with total nitrogen levels.

Yoshida, Mitsuhiro et al. "Ecological dynamics of the toxic bloom-forming cyanobacterium Microcystis aeruginosa and its cyanophages in freshwater."

A study by Yoshida et al. sought to investigate the role of cyanophages on M. aeruginosa communities in samples isolated from Lake Mikata (Japan) (21). The same site was sampled each month from April to October, 2006. To quantify total M. aeruginosa, a qPCR amplification of the phycocyanin intergenic spacer (PC-IGS) was used (present in all strains of Microcystis aeruginosa). A second qPCR assay was used to quantitatively detect potentially infectious Ma-LMM01 cyanophages using 36 the primers targeting the viral sheath protein-encoding gene (g91). The study also used qPCR to quantify the proportion of one of the components of the microcystin synthetase gene (mcyA) to assess the abundance of toxic strains of Microcystis aeruginosa. The results of the study indicated that as cyanophage abundance increased, there was a temporal decline in total Microcystis aeruginosa abundance.

The study also noted that mcyA producing strains had a greater abundance in April-

August (>18%) than in September-November (0.50 to 2.25%). The decline in mcyA strains coincided with a rise and dominance in non-toxic strains when the bloom reemerged. The data suggests that cyanophages may have induced the decline in the microcystin-producing subpopulation, which allowed for the shift to non-toxic strain dominance in the early fall months (September-November).

Further Support for Cyanophage-Mediated Genotypic Succession of Microcystis aeruginosa over a Bloom Period

Similar to the findings of Yoshida et al., emerging research by Driscoll et al. from the Dreher Lab at Oregon State University has proposed that a Microcystis infecting virus Ma-LMM01 strain may be responsible for bloom collapse observed in

San Francisco Delta watershed (42). This strain of Ma-LMM01 is genetically similar to the Ma-LMM01 strain that was first isolated by Yoshida et al. from 2006 samples taken from Lake Mikata, Japan (21). Bloom collapse may provide an ecological niche for the rise of another bloom-forming cyanobacterial species. This cyanobacterial species could subsequently establish dominance if it has some type of innate resistance to the factor that caused the demise of the previous population. Besides the research publication by Yoshida et al., the topic of Ma-LMM01 mediated bloom 37 collapse is not widely described in research literature (21). The relationship between Ma-LMM01 and bloom population dynamics is an active area of research in the Dreher lab. In this study, samples collected for twelve weeks in 2011 and 2012 were analyzed with quantitative PCR to quantify populations from each testing site.

Samples from the Mildred Island site showed a rapid decline in Microcystis abundance. Metagenome sequencing of the DNA extracted before and after the bloom collapse identified twelve putative phage genomes that were actively infecting cells within the samples. One of these genomes strongly resembled virus

Ma-LMM01. Further investigation showed that the reduction in the quantity of the

Ma-LMM01-like virus correlated with the reduction in total Microcystis cells after the bloom demise. This research may shine light on the occurrence of virus- mediated bloom collapse and investigate a possible mechanism for genotypic succession in Microcystis aeruginosa blooms.

CONCLUSION

Rising global temperatures, along with anthropogenic sources of aquatic pollution have led to the eutrophication of many bodies of water across the planet.

As a result of this, it is predicted that the extent and severity of cyanobacterial blooms is set to increase, potentially affecting human health, recreation, and watershed condition. This study intended to investigate the response of toxic and non-toxic strains of Microcystis aeruginosa to increased oxidative conditions induced by the incidence of high light over a 14 day growth period. The results indicate that UTEX 2667 and CPCC 299 (toxic strains) of Microcystis aeruginosa were less inhibited by exposure to 300 μmol of photons m−2 s−1 (12 light / 12 hour dark cycle) than were PCC 7005 and UTEX 2386 (non-toxic strains). This result is in keeping with evidence that microcystin may play a role in stabilizing photosynthesis enzymes such as RuBisCO, which was reported by Zilliges et al. (8). This study also adapted a scopoletin assay to measure the amount of hydrogen peroxide produced from exposure to high light, which is caused by the over-excitation of electron transport machinery in a mechanism detailed by Kozuleva et al. (7). The results of

Day 11 and Day 14 scopoletin measurements indicate that on both days, the high light treatments of all samples produced a quantifiable hydrogen peroxide increase in the range of 10-9 nM. These results are consistent with findings that the cause of stress induced by high light on cyanobacteria is oxidative in nature.

The study included an investigation of published scientific findings on 2-cys peroxiredoxin, an enzyme that has been studied in related species as one that helps the organism “cope” with increased levels of peroxide concentrations. From this 39 investigation, it appears that further investigation into 2-cys peroxiredoxin expression by Microcystis aeruginosa under oxidative conditions may be fruitful.

Lastly the study looked to investigate the causes of genotypic variation from toxic to non-toxic strains observed in environmental studies focused on Microcystis aeruginosa dominated blooms. Several different mechanisms have been proposed.

These mechanisms range from cyanophages, microcystin-degrading bacteria, nutrient limitation/meszooplankton grazing, amoebae grazing, to the changes in light intensity during a bloom season that causes vertical genotypic variation. It appears that no clear consensus has been reached to explain this phenomenon. This study demonstrated that toxic and non-toxic strains of Microcystis aeruginosa respond differentially to high light, which provides support to the hypothesis that light intensity could play a role in this observed genotypic succession. There is also ongoing research on the toxic bloom collapse mediated by viral (cyanophage) infection, which may provide further insight into this phenomenon.

The findings of this study provide a basis for the further assessment of how high light / oxidative conditions affect toxic and non-toxic Microcystis aeruginosa. As a next step, it would be beneficial to use qPCR to assess the expression of 2-cys peroxiredoxin under a similar experimental design used in this study. It would also be advantageous to investigate whether Microcystis aeruginosa can respond to oxidative stress in the presence of a gene knock out for 2-cys prx. In total, further understanding of factors that favor the growth/inhibition of cyanobacteria and

Microcystis aeruginosa-dominated blooms may lead to the development of potential management strategies for this pressing environmental concern. 40

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