Comparative!GrowthStudyofToxicandNon :Toxic Microcystis ...

1

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

by

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

2

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.

3

Key Words: Microcystis aeruginosa, oxidative stress, cyanobacterial blooms, scopoletin assay, microcystin, photooxidative stress, toxic strain dominance Corresponding e-mail address: [email protected]

4

Copyright by Neil Rajput May 15, 2014

All Rights Reserved

5

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

6

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.

7

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 of Peroxiredoxin Genes

16

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

17

RESULTS AND DISCUSSION

18

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

18

H2O2 Concentration Determination using Adapted Scopoletin Assay 22 Investigation of 2-‐cys Peroxiredoxin 27

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

29

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 of Microcystis aeruginosa over a Bloom Period

36

CONCLUSION 38 REFERENCES 40

8

LIST OF FIGURES

Figure

Figure 1. Structure of Microcystin-‐LR. Microcystin (mcy) gene cluster 4

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

6

Figure 3. Enlarged Grid of Hemocytometer 16

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

20

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-‐Toxic"

21

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 Complete Genomes of Microcystis Aeruginosa Strains

28

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.

30

9

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 (nM) -‐ Growth Day 11

25

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

25

Table 5. Amino Acid Sequence of 2-‐cys Peroxiredoxin (BAS1) 27

LIST OF EQUATIONS

Equation

Equation 1. Cell Density Equation for Hemocytometer Counting 15

10

DEDICATION

This thesis is dedicated to the professors and faculty

of the Microbiology department at Oregon State University

for their expertise and continual support.

1

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).

9

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.

10

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.

11

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.

17

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.

18

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.

20

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

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

0.00E+00 5.00E+06 1.00E+07 1.50E+07 2.00E+07 2.50E+07 3.00E+07 3.50E+07 4.00E+07

0 2 4 6 8 10 12 14 16

Cell Density (cells/ml)

Day

Microcystis aeruginosa Strain PCC 7005 - "Non-Toxic"

LL HL

0.00E+00 5.00E+06 1.00E+07 1.50E+07 2.00E+07 2.50E+07 3.00E+07 3.50E+07 4.00E+07

0 2 4 6 8 10 12 14 16


Day

Microcystis aeruginosa Strain UTEX 2667 - "Toxic"

LL HL

21

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

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

0.00E+00 5.00E+06 1.00E+07 1.50E+07 2.00E+07 2.50E+07 3.00E+07 3.50E+07 4.00E+07

0 2 4 6 8 10 12 14 16


Day

Microcystis aeruginosa Strain CPCC 299 - "Toxic"

LL HL

0.00E+00 5.00E+06 1.00E+07 1.50E+07 2.00E+07 2.50E+07 3.00E+07 3.50E+07 4.00E+07

0 2 4 6 8 10 12 14 16


Day

Microcystis aeruginosa Strain UTEX 2386 - "Non-Toxic"

LL HL

22

H2O2 Concentration Determination using Adapted Scopoletin Assay

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

Sample Fluorescence

Experimentally Derived

Concentrations of H2O2 (nM)

Catalase-‐treated Water (Control)

20504

BG-‐11 (Media) 19932

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 HL/S 18103 281.69

PCC 7005 LL 19168 148.24 PCC 7005 LL/S 18240 264.53

UTEX 2667 HL 19108 155.76

UTEX 2667 LL 19309 130.57

CPCC 299 HL 19199 144.36 CPCC 299 LL 19403 118.79 UTEX 2386 HL 19202 143.98

UTEX 2386 LL 19337 127.06

*average background fluorescence subtracted

23

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

Sample Fluorescence

Experimentally Derived

Concentrations of H2O2 (nM)

Catalase-‐treated Water (Control)

20104

BG-‐11 (Media) 19787

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 HL/S 18374 212.82

PCC 7005 LL 19377 85.84 PCC 7005 LL/S 18689 172.94

UTEX 2667 HL 19200 108.25

UTEX 2667 LL 19433 78.75

CPCC 299 HL 18855 151.92 CPCC 299 LL 19116 118.88 UTEX 2386 HL 18795 159.52

UTEX 2386 LL 19082 123.18

*average background fluorescence subtracted

24

Figure 8. Scopoletin Assay Standard Curve - Growth Day 11

Figure 9. Scopoletin Assay Standard Curve - Growth Day 14

y = -‐7.8987x + 20055

0

5000

10000

15000

20000

25000

0 500 1000 1500 2000 2500

Units Fluorescence

H2O2 Concentration (nM)

Scopoletin Assay Standard Curve - Growth Day 11

y = -‐7.9803x + 20351

0

5000

10000

15000

20000

25000

0 500 1000 1500 2000 2500

Units Fluorescence

H2O2 Concentration (nM)

Scopoletin Assay Standard Curve - Growth Day 14

25

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

Strain Difference in H2O2

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

Strain Difference in H2O2

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.

27

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

28

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.

38

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

REFERENCES

1. Dziallas, Claudia, and Hans-Peter Grossart. 2011. Increasing oxygen radicals and water temperature select for toxic Microcystis sp. PloS one. 6:e25569.

2. Vézie, C., et al. 2002. Effect of nitrogen and phosphorus on growth of toxic

and nontoxic Microcystis strains and on intracellular microcystin concentrations." Microbial ecology. 43:443-‐454.

3. Department of Health and Human Services CDC. 2010. Harmful Algal

Blooms: Environmental Hazards and Health Effects. Retrieved from US Center for Disease Control: http://www.cdc.gov/hab/

4. Paerl, Hans W., Nathan S. Hall, and Elizabeth S. Calandrino. 2011.

Controlling harmful cyanobacterial blooms in a world experiencing anthropogenic and climatic-‐induced change. Science of the Total Environment. 409:1739-‐1745.

5. Stewart, Ian, and Ian R. Falconer. 2008. Cyanobacteria and cyanobacterial

toxins. Oceans and human health: risks and remedies from the seas. 271-‐296.

6. McFadden, Geoffrey I., and Giel G. van Dooren. 2004. Evolution: red algal genome affirms a common origin of all plastids. Current Biology. 14:514-‐516.

7. Kozuleva, Marina A., et al. 2012. Photosynthetic electron flow to oxygen

and diffusion of hydrogen peroxide through the chloroplast envelope via aquaporins. Biochimica et Biophysica Acta (BBA)-‐Bioenergetics. 1817:1314-‐1321.

8. Zilliges, Yvonne, et al. 2011. The cyanobacterial hepatotoxin microcystin

binds to proteins and increases the fitness of Microcystis under oxidative stress conditions. PloS one. 6:e17615.

9. Wiedner, Claudia, et al. 2003. Effects of light on the microcystin content of

Microcystis strain PCC 7806. Applied and Environmental Microbiology. 69:1475-‐1481.

10. UTEX The Culture Collection of Algae. UTEX The Culture Collection of

Algae. N.p., n.d. Web. http://web.biosci.utexas.edu/utex/mediaDetail.aspx?mediaID=26.

11. Misson, Benjamin, and Delphine Latour. 2013. Vertical Heterogeneity of

Genotypic Structure and Toxic Potential within Populations of the Harmful Cyanobacterium Microcystis aeruginosa. Advances in Microbiology. 3: 27.

41

12. Penn, Kevin, et al. 2014. Secondary metabolite gene expression and interplay of bacterial functions in a tropical freshwater cyanobacterial bloom. The ISME journal.

13. Sielaff, Heike, et al. 2003. The mcyF gene of the microcystin biosynthetic

gene cluster from Microcystis aeruginosa encodes an aspartate racemase. Biochem. J. 373: 909-‐916.

14. Watanabe, Mariyo F., et al. 1995. Toxic Microcystis. CRC press.

15. Klein, Annaleise R., Darren S. Baldwin, and Ewen Silvester. 2013. Proton

and Iron Binding by the Cyanobacterial Toxin Microcystin-‐LR." Environmental science & technology. 47:5178-‐5184.

16. Leunert, Franziska, et al. 2014. Phytoplankton response to UV-‐generated

hydrogen peroxide from natural organic matter. Journal of Plankton Research. 36:185-‐197.

17. Corbett, Jean T. 1989. The scopoletin assay for hydrogen peroxide A review

and a better method. Journal of biochemical and biophysical methods 18:297-‐307.

18. Counting Cells with a Hemocytometer - Protocol. Counting Cells with a

Hemocytometer -‐ Protocol. N.p., n.d. Web. http://web.mnstate.edu/provost/CountingCellsHemocytometer.pdf.

19. Yang, Chen, et al. 2013. Whole-‐genome sequence of Microcystis aeruginosa

TAIHU98, a nontoxic bloom-‐forming strain isolated from Taihu Lake, China." Genome announcements. 1:e00333-‐13.

20. Cooper, William J., et al. 1988. Photochemical formation of hydrogen

peroxide in natural waters exposed to sunlight. Environmental science & technology. 22:1156-‐1160.

21. Yoshida, Mitsuhiro, et al. 2008. Ecological dynamics of the toxic bloom-‐

forming cyanobacterium Microcystis aeruginosa and its cyanophages in freshwater. Applied and environmental microbiology. 74:3269-‐3273.

22. Kardinaal, W. Edwin A., et al. 2007. Microcystis genotype succession in

relation to microcystin concentrations in freshwater lakes. Aquatic Microbial Ecology. 48:1-‐12.

23. Zhu, Lin, et al. 2014. Ecological dynamics of toxic Microcystis spp. and

microcystin-‐degrading bacteria in Dianchi Lake, China. Applied and environmental microbiology. AEM-‐02972.

42

24. Latifi, Amel, Marion Ruiz, and Cheng‐Cai Zhang. 2009. Oxidative stress in cyanobacteria. FEMS microbiology reviews. 33:258-‐278.

25. Yamamoto H, Miyake C, Dietz KJ, Tomizawa K, Murata N & Yokota A.

1999. Thioredoxin peroxidase in the cyanobacterium Synechocytis sp. PCC 6803. FEBS Lett. 447:269–273.

26. Perelman A, Uzan A, Hacohen D & Schwarz R. 2003. Oxidative stress in

Synechococcus sp. strain PCC 7942: various mechanisms for H2O2 detoxification with different physiological roles. J Bacteriol. 185:3654–3660.

27. Hall, Andrea, P. Andrew Karplus, and Leslie B. Poole. 2009. Typical 2-‐Cys

peroxiredoxins–structures, mechanisms and functions. FEBS journal. 276:2469-‐2477.

28. Rantala, Anne, et al. 2004. Phylogenetic evidence for the early evolution of

microcystin synthesis. Proceedings of the National Academy of Sciences of the United States of America. 101:568-‐573.

29. Rohrlack, Thomas, et al. 2001. Effects of cell-‐bound microcystins on

survival and feeding of Daphnia spp. Applied and environmental microbiology. 67:3523-‐3529.

30. Schatz, Daniella, et al. 2007. Towards clarification of the biological role of

microcystins, a family of cyanobacterial toxins." Environmental microbiology. 9:965-‐970.

31. Börner, Thomas, and Elke Dittmann. 2005. Molecular biology of

cyanobacterial toxins. Harmful cyanobacteria. Springer. Netherlands. 25-‐40.

32. Mostofa, Khan MG, et al. 2013. Photoinduced Generation of Hydroxyl Radical in Natural Waters." Photobiogeochemistry of Organic Matter. Springer. Berlin Heidelberg. 209-‐272.

33. Kieber, Robert J., and G. R. Helz. 1986. Two-‐method verification of

hydrogen peroxide determinations in natural waters. Analytical chemistry 58:2312-‐2315.

34. Dons, Lone E., et al. 2013. Role of the Listeria monocytogenes 2-‐Cys

peroxiredoxin homologue in protection against oxidative and nitrosative stress and in virulence. Pathogens and disease.

35. Cui, Hongli, et al. 2012. Genome-‐wide analysis of putative peroxiredoxin in

unicellular and filamentous cyanobacteria. BMC evolutionary biology. 12: 220.

43

36. Li, Daming, et al. 2012. Quantification of microcystin-‐producing and non-‐microcystin producing Microcystis populations during the 2009 and 2010 blooms in Lake Taihu using quantitative real-‐time PCR." Journal of Environmental Sciences. 24:84-‐290.

37. Van Wichelen, Jeroen, et al. 2010. Strong effects of amoebae grazing on the

biomass and genetic structure of a Microcystis bloom (Cyanobacteria). Environmental microbiology. 12:2797-‐2813.

38. Yoshida, Mitsuhiro, et al. 2007. Dynamics of microcystin-‐producing and

non-‐microcystin-‐producing Microcystis populations is correlated with nitrate concentration in a Japanese lake. FEMS microbiology letters. 266:49-‐53.

39. Joung, Seung-Hyun, et al. 2011. Correlations between environmental

factors and toxic and non-‐toxic Microcystis dynamics during bloom in Daechung Reservoir, Korea. Harmful Algae. 10:188-‐193.

40. Gobler, C. J., et al. 2007. Interactive influences of nutrient loading,

zooplankton grazing, and microcystin synthetase gene expression on cyanobacterial bloom dynamics in a eutrophic New York lake. Harmful Algae. 6:119-‐133.

41. Bozarth, Connie S., et al. 2010. Population turnover in a Microcystis bloom

results in predominantly nontoxigenic variants late in the season. Applied and environmental microbiology. 76:5207-‐5213.

42. Connor, Driscoll. Investigating the role of viruses in Microcystis sp. bloom

collapse. WALPA RSS. Washington State Lake Protection Association, n.d. Web. http://www.walpa.org/investigating-‐the-‐role-‐of-‐viruses-‐in-‐Microcystis-‐sp-‐bloom-‐collapse.

43. Murata, Norio, et al. 2007. Photoinhibition of photosystem II under

environmental stress." Biochimica et Biophysica Acta (BBA)-‐Bioenergetics. 1767:414-‐421.

44. Ding, Yi, et al. 2012. Hydrogen peroxide induces apoptotic-‐like cell death in

Microcystis aeruginosa (Chroococcales, Cyanobacteria) in a dose-‐dependent manner. Phycologia. 51:567-‐575.

45. Campos, Alexandre, and Vitor Vasconcelos. 2010. Molecular mechanisms

of microcystin toxicity in animal cells. International journal of molecular sciences. 11:268-‐287.

44

46. Pereira, Susana R., Vítor M. Vasconcelos, and Agostinho Antunes. 2013. Computational study of the covalent bonding of microcystins to cysteine residues–a reaction involved in the inhibition of the PPP family of protein phosphatases. FEBS Journal. 280:674-‐680.

47. Pasternak, Zohar, et al. 2012. By their genes ye shall know them: genomic

signatures of predatory bacteria. The ISME journal. 7:756-‐769.

Comparative!GrowthStudyofToxicandNon :Toxic Microcystis ...

Documents

Transcript of Comparative!GrowthStudyofToxicandNon :Toxic Microcystis ...