Assessment of Biodegradation and Toxicological Effects Of

Home , Microbial biodegradation

ASSESSMENT OF BIODEGRADATION AND TOXICOLOGICAL EFFECTS OF

MICROCYSTINS

A dissertation submitted

to Kent State University in partial

fulfillment of the requirements for the

degree of Doctor of Philosophy

Anjali Krishnan

August, 2019

Except for previously published materials

Dissertation written by

Anjali Krishnan

B.Sc., University of Mumbai, 2011

M.Sc., University of Mumbai, 2013

Ph.D., Kent State University, 2019

Approved by

Xiaozhen Mou, Ph.D., Chair, Doctoral Dissertation Committee

Gary Koski, Ph.D., Members, Doctoral Dissertation Committee

Darren Bade, Ph.D.

Joseph Ortiz, Ph.D.

Michael Strickland, Ph.D.

Accepted by

Dr. Ernest Freeman, Ph.D., Chair, Department of Biological Sciences

Dr. James L. Blank, Ph.D., Dean, College of Arts and Sciences

TABLE OF CONTENTS ...... iii LIST OF FIGURES ...... v LIST OF TABLES ...... viii ACKNOWLEDGEMENTS ...... ix I.GENERAL INTRODUCTION AND OVERVIEW ...... 1 REVIEW OF LITERATURE ...... 1 REFERENCES ...... 22 II. IDENTIFICATION AND CHARACTERIZATION OF MICROCYSTIN DEGRADING BACTERIA FROM LAKE ERIE...... 38 PREFACE ...... 38 ABSTRACT ...... 38 INTRODUCTION ...... 39 METHODS ...... 41 RESULTS ...... 44 DISCUSSION ...... 44 ACKNOWLEDGEMENTS ...... 48 REFERENCES ...... 54 III TAXONOMIC AND FUNCTIONAL HETEROGENEITY OF BACTERIOPLANKTON IN LAKE ERIE...... 59 PREFACE ...... 59 ABSTRACT ...... 59 INTRODUCTION ...... 62 METHODS ...... 65 RESULTS ...... 69 DISCUSSION ...... 72

iii

REFERENCES ...... 87 IV. IDENTIFICATION OF NOVEL MICROCYSTIN DEGRADING PATHWAY ADOPTED BY BACTERIOPLANKTON FROM LAKE ERIE USING TRANSPOSON MUTAGENESIS AND LC-MS...... 95 ABSTRACT ...... 95 INTRODUCTION ...... 96 METHODS ...... 99 RESULTS ...... 103 DISCUSSION ...... 105 CONCLUSION ...... 107 REFERENCES ...... 114 V.CHARACTERIZATION OF MICROCYSTIN-INDUCED APOPTOSIS IN HepG2

HEPATOMA CELLS...... 117

ABSTRACT ...... 117 INTRODUCTION ...... 118 METHODS ...... 120 RESULTS ...... 123 DISCUSSION ...... 123 CONCLUSION ...... …. 127 ACKNOWLEDGEMENTS...... 127 REFERNCES ...... 133 VI.SUMMARY CHAPTER ...... 139 GENERAL DISCUSSION ...... 139 GENERAL CONCLUSION ...... 144 REFERENCES ...... 145 APPENDIX ...... 147

LIST OF FIGURES

Figure 1. Cyanobacterial blooms as seen in western basin of Lake Erie in the Summer of 2016.19

Figure 2. Schematic representation of mlr and xenobiotic degradation pathway ...... 20

Figure 3. Schematic representation of the extrinsic and intrinsic apoptotic pathway ...... …..21

Figure 4. Changes of MC-LR concentration (left axis, solid gray line) and optical density OD600

(right axis, dash black line) in BG11 growth media supplied with 1 µg/ml of MC-LR and bacterial isolates...... 50

Figure 5. A neighbor-joining phylogenetic tree based on partial sequence of 16S rRNA gene showing the relationship sequences of isolated MC-degrading bacteria and their closely related relatives. Gray circles are to label previously identified MC-degrading bacteria. The GenBank accession numbers of the sequences are shown in parentheses. Bootstrap values that are higher than 50% are show at the branch nodes (1,000 resampling). The scale bar represents 0.02 nucleotide substitutions per position ...... 51

Figure 6. Effect of temperature on MC-LR degradation by MC- degrading bacteria...... 52

Figure 7. Effects of pH on degradation of MC-LR by MC-degrading bacteria...... 53

Figure 8. Microcystin degradation of the MC+ isolates as seen in the BIOLOG MT2 screening.

Bacterial isolates which showed a significant increase in their optical density for MC-LR concentrations of 0.1, 1 and 10µg/ml were depicted in this bar graph ...... 79

Figure 9. Microscopic observation for gram staining performed for MC-degrading bacterial isolates with morphologies of bacilli (a) ,cocci (b) and coccobacilli (c) ...... 80

Figure 10. A neighbor-joining phylogenetic tree based on partial sequence of 16S rRNA gene showing the relationship sequences of isolated MC-degrading bacteria and their closely related

relatives. The GenBank accession numbers of the sequences are shown in parentheses. Bootstrap values that are higher than 50% are show at the branch nodes (1,000 resampling). The scale bar represents 0.02 nucleotide substitutions per position...... 81

Figure 11: Gel image for the mlrA based amplification performed for MC+ isolates. mlrA amplification was observed for the positive control Sphingosinicella microcystinivorans (16998) and Paucibacter toxinivorans (19791) ...... 82

Figure 12: Effect of temperature on MC-LR degradation rate by MC-degrading bacteria as shown in a heat map. The gradient of colors from blue to red indicate the degradation rate for the MC+ isolates ...... 83

Figure 13: Effect of pH on MC-LR degradation rate by MC-degrading bacteria as shown in a heat map for MC+ isolates. The gradient of color from blue to red indicate the degradation rates for the

MC+ isolates ...... 84

Figure 14: Heat map representing the degradation rates (µg/ml/hr) for isolates when incubated in a suspension containing other organic nutrients. The gradient of colors from blue to red represent the degradation rates of the isolates. Control represents the degradation rate of the isolates when incubated in a suspension containing only MC-LR and test represents the microcystin degradation rate for the isolates when incubated in suspension containing MC-LR and organic C source. . . .85

Figure 15: MC-LR degradation rates (µg/ml/hr) as measured for mixed culture MC+ isolates.

Degradation rates were compared between the individual MC+ isolates and when in mixture. * has been added for the isolate combinations which have a higher degradation rates than the individual isolates...... ………………………………………………………………… .86

Figure 16. MC-LR degradation pathway: The figure demonstrates the mlr based degradative pathway and the potential xenobiotic metabolic degradative pathway (Mou et al.,

2013). ………...... 108

Figure 16: MC-LR degradation of the Tn5 mutants was identified using absorbance measurements in BIOLOG MT2 screening.………………………………………………………………… …109

Figure 17. Changes in optical density (Y axis) over time (X-axis) for the growth curve of LEw-24 and its mutants A45, F87, F72, D1 and F92. ……………………………………………………111

Figure 18. HPLC/MS figures for the identification of the degradation products and intermediates.112

Figure 19. Effects of various concentrations of MC-LR on metabolic activity of HepG2 cells with optical density values before (0 hr) and after (10 hr) incubation with Alamar Blue...... 129

Figure 20. Confocal microscopy pictures of control (untreated) (a) and 5µM MC-LR treated (b)

HpeG2 cells after 24 hrs of incubation. . . ………………………………………………………130

Figure 21. Histogram showing fluorescence for untreated HepG2 cells (black) grown in DMEM medium and MC-LR treated (blue) HepG2 cells stained with caspases 3/7 specific binding dye

(green) (a), caspases 8 specific stain (red) (b) and caspases 9 FITC stain (blue) (c) ...... 131

Figure 22. Histogram representing the fluorescence for treated, caspase-8 (red, a) and caspase-9 expressing (blue, b) HepG2 cells, observed at different time points of 0, 4, 8, 12 and 24 hrs of MC-

LR treatment. Control (black, untreated) HepG2 cells are also shown in this figure ...... 132

vii

LIST OF TABLES

Table 1. Highest microcystin concentration recorded for various lakes around the world …. .17

Table 2. Isolated microcystin degrading bacteria ...... 18

Table 3. Phenotypic characteristics of microcystin-degrading bacterial isolates obtained from water samples of Lake Erie ...... 49

Table 4. Morphological characteristics of microcystin degrading bacteria ...... 76

Table 5. Combination of the MC-degrading isolates used for mixed culture degradation . . . .78

Table 6. Sequence data showing the BLAST match for the sequences ...... 110

Appendix

Table 7. Pure culture isolates from samples collected from a biofilter in Toledo, Akron (water and sediment), Ravenna (water and sludge), Alliance WTP (backwash, sludge) and Sandusky (water) during the summer of 2015 and 2016 ...... 147

Table 8. Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Akron water treatment plant in the summer of 2018...... 153

Table 9. Pure culture isolates and their colony characteristics as seen on R2A agar from the water and sludge samples collected from the Alliance water treatment plant in the summer of 2018. .155

Table 10. : Pure culture isolates and their colony characteristics as seen on R2A agar from the water and sludge samples collected from the Ravenna water treatment plant in the summer of 2018.

...... ……………………………………………………….159

Table 11. Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Kent water treatment plant in the summer of 2018...... 165

Table 12. Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Sandusky water treatment plant in the summer of 2018...... 169 viii

Acknowledgements

At this moment, I would like to thank and acknowledge all who have helped me during my long and arduous academic journey. First and foremost, I would like to express deep gratitude for my

PhD advisor Dr. Xiaozhen Mou. She has been a constant source of motivation and encouragement which has led me to successfully complete my experiments and write my dissertation. She helped me identify my errors and rectify them, which has enabled me to become the educator/ scientist that I am today. She has always been supportive of my ideas and helped me progress in the right direction. Apart from being a mentor, she has also become my friend and has always believed in me. I will forever be grateful and cherish the time that I got to spend with her.

I would like to extend my deep felt gratitude to all my committee member, Dr. Gary Koski,

Dr. Darren Bade and Dr. Joseph Ortiz for their constant support and help over the past years. I would like to thank National Science Foundation, Environmental Protection Agency, Ohio Lake

Erie Protection Fund, and Kent State University Department of Biological Sciences for providing me with the required funding and support for successful completion of my experiments

I would like to thank my friends in the Mou and Blackwood lab; Anna Droz and Andrew

Eagar for their friendship and fun during trying times. I would also like to thank my friends Tharani

Manickavasagam, Anurag Jamaiyar, Jayakrishnan Ajaykumar, Vinay Joshi and Shelly Fisher for their help, constant advice, friendship and support.

I would like to dedicate this dissertation and thank my parents N.A Krishnan and Vasantha

Krishnan for their constant support and believing in me during my time at Kent State University.

I would also like to thank my sister Aditi Krishnan and my brother-in-law Vivek Raju for always believing in me and providing with me emotional support during my time at Kent State University.

Chapter 1 General Introduction and Overview

Preface

The first chapter for my dissertation is being made into a manuscript as a review paper for

publication in Toxins journal. My co-author and dissertation adviser Xiaozhen Mou, has helped

me significantly with the writing of the paper.

Introduction

MCs (MCs) are a class of liver toxins to human and animals alike (Labine et al., 2009). MCs

are produced as secondary metabolites by a number of freshwater cyanobacteria, including

Microcystis, Planktothrix, Anabaena and Oscillatoria genera. During the growth season of

cyanobacteria (Fig 1), MCs are often at high concentrations (100-2000 µg/L) in freshwater

environments (Table 1) (WHO guideline concentration, 1 µg/L)which possess a great health

concern when these waters are used as drinking or recreation sources (Sedmak et al., 2018). A

vast number of studies have focused on MC synthesis and their regulation factors in natural

environment (Jahnichen et al., 2011). However, less is known on biological degradation and

hepatotoxic effects of these toxins to human, which were the foci of this dissertation research.

1.1 Structure of MCs

MCs possess a cyclic heptapeptide structure. This structure involves the presence of seven

amino acids which are connected by peptide bonds in a cyclic structure (Rinehart et al., 1988).

This cyclic configuration of MCs has a number of variants due to the replacement of the α-amino

acids in the cyclic structure. Over 90 isoforms of MCs have been identified based on the variation of the α-amino acid in the structure (Pearson et al., 2003). A few variants of MCs, based on the variation in their structure are, MC-LR, MC-RR and MC-LF, MC-YR; of which, MC-LR is the most prevalent and toxic (Dietrich et al., 2005). In MC-LR, we have leucine (L) and arginine (R) in these positions for α-amino acids (Fontanillo et al., 2018) which makes MC-LR more toxic than the other MC variants identified, due to easier uptake of this variant by the bile acid transport system (Schmidt et al., 2014). A unique feature of MC-LR is the presence of the C20 amino acid

(3-amino-9-methoxy-2, 6, 8-trimethyl-10-phenyldeca-4, 6-dienoic acid) which is also known as

Adda (Rinehart et al., 1988). This Adda chain is consistent among the MC- variants and can be used to quantify MCs. MC-LR also shows the presence of a unique amino acid MdhA (methyl dehydroalanine). Both Adda and MdhA are crucial for binding of MCs to protein phosphatases in organism.

1.2 Production of MCs and its regulating factors

Studies have identified that mcy genes are responsible for MC production in Microcystis aruginosa and other cyanobacteria such as Anabaena and Planktothrix species (Tillet et al., 2000).

The gene cluster mcy possess 55kbp of DNA which encodes for mcyA-mcyJ. These genes encode

48 catalytic reactions that are required for MC synthesis (Tillet et al., 2000). The primary

precursors involved in the MC synthesis are phenyl acetate, malonyl-CoA, SAM, glutamate, serine, alanine, leucine, D-methyl-iso-aspartate, and arginine (Tillet et al., 2000; Dittmann et al.,

1997). The Adda ring is formed by mcyG, mcyD and mcyE cluster of genes. After the formation of the Adda ring the final step involves condensation reaction catalyzed by the carboxy terminal in McyC.

A number of studies have found that environmental factors like irradiance, Pi and Fe3+ play significant role in controlling MC production (Jahnichen et al., 2007, 2011; Dai et al., 2016) with increased MC production seen as a result of lower irradiance and lower Pi supply increased supply of Fe3+. Some of the other environmental factors which affect MC production in cyanobacteria are nitrogen content (Orr et al., 1998; Ginn et al., 2010), sulphur and phosphorous (Oh et al., 2000;

Juliana et al., 2019), and oxygen saturation (Wicks and Thiel, 1990, Kotak et al., 2000; Srivastava et al., 2012). A positive correlation was observed with increasing concentrations of total phosphorous concentrations in the bloom affected lake (Rinta-Kanto et al., 2009). Solar radiation and oxygen saturation are also shown to be positively correlated to MC production by cyanobacteria (Wicks and Thiel, 1990). Increased sulfur has also been found to increase MC production by cyanobacteria (Jahnichen et al., 2011). An alkaline pH of 9.5 and a temperature of

20ºC has been found to yield maximum production of MC (Van der Westhuizen and Eloff, 1985).

More than 90 variants of MC have been observed during blooms (Vesterkvist et al., 2012).

The factors affecting the production of the different MC congeners in Microcystis aeruginosa HUB 5-2-4 are light intensity and nutrient supply (Hesse and Kohl, 2012). In some 3

other species like Planktothrix agardhii, the ratio of production for MC-LR and MC-RR was found to be affected by photo irradiance and availability of amino acids, mainly leucine and arginine (Tonk et al., 2005). Studies have been performed to analyze MC congener production under different environmental conditions (Xie et al., 2016), where light and temperature did not affect the production of the toxin but factors like nitrogen and phosphorous were found to affect the toxin production.

1.3 MC degradation

1.3.1 Physicochemical degradation

MCs are sensitive to light and complete photo degradation of MC occurs within 5 days at

365nm where MC-LR is degraded at the ADDA ring (Schmidt et al., 2014). MCs are also resistant to physicochemical and biological stresses such as temperature, pH and certain proteases (Harada,

1996; Tsuji et al., 1994 and Jones et al., 1994).

1.3.2 Biological degradation

MC- degrading bacteria have also been isolated from sources such as drinking water sludge and the degradation rate measured using anoxic conditions (Mat et al., 2014). A number of other studies have examined the potential of these MC degrading bacteria in biological filters as well

(Shimizu et al., 2013). In these filters complete removal of the toxin was found to occur within 4 days with a lag period of three days (Wang et al., 2007). Therefore, recent culture-dependent and independent studies consistently suggest that MC-degrading bacteria might be more taxonomic diverse than previously thought (Table 2). 4

Naturally occurring bacteria has been to shown to possess the ability to degrade MC with increasing number of studies reporting the presence of MC- degrading bacteria in lakes affected with cyanobacterial blooms (Eleuterio and Batista, 2010). Culture independent work for MC degradation has shown that bacterial communities present in lakes exposed to cyanobacterial blooms, can degrade MC (Christoffersen et al., 2002; Mou et al., 2013). A 2-day lag phase has been observed for the degradation of MCs using bacterial communities present in such lakes. mlr gene based pathway

The degradative pathway adopted by MC-degrading bacteria has been firstly identified in

Sphingomonas sp. ACM-3962 as an mlr gene based step-wise cleavage process (Jones et al., 1994).

Enzyme MlrA (microcystinase) initiate the degradation by breaking up the ring and linearizing the cyclic MC-LR. The linearized MC-LR has much lower toxicity than cyclic MC-LR (Jones et al.,

1994). Next, a serine peptidase (MlrB) catalyzes the linearized MC-LR at the Ala-Leu peptide bond and produces a tetrapeptide (Jones et al., 1994). Finally, the third enzyme (MlrC), cuts the peptide bonds randomly resulting in undetectable peptide fragments and amino acids. However,

MlrB has been shown to lose it activity in some strains isolated from Lake Taihu (Jiang et al.,

2011) with high activity observed for MlrA expression. Since the initial cleavage is the most critical step in MC-LR degradation, so far genetic analysis of MC-LR degradation is mostly based on analysis of mlrA genes (Jones et al., 1994).

non-mlr pathway

The presence of mlrA has been universally identified in MC-degrading strains that belonged to the Sphingomonadaeceae family within the genera of Sphingosinicella, Sphingopyxis,

Sphingomonas and Novosphingobium (Kormas et al., 2013) However one MC-degrading

Actinobacteria showed the absence of the mlrA based on PCR assay (Manage et al., 2010).

Following this study there has been multiple reports on the absence of mlr gene in MC- degrading bacterial isolates (Yang et al., 2014; Kansole et al., 2016). This suggests two possibilities: 1) PCR programs that were designed for amplifying mlrA genes in Sphinogomonas and related taxa are not applicable to taxonomically distinct lineages and 2) this actinobacteria has an alternative MC-

LR degradative pathway.

A recent metagenomic study supported the second possibility (Mou et al., 2013).

Metagenomes of Lake Erie bacterioplankton contained extremely low occurrence of mlr genes, and the abundance of these mlr genes showed no significant changes in response to MC-LR addition. On the other hand, xenobiotic metabolism related genes were largely present in the MC metagenome (Fig 2).

Xenobiotic metabolism is widely spread among the different domains of life (Jannsen et al., 2005) and has shown its role in MC degradation in aquatic eukaryotes (Pergrin-Alvarez et al.,

2009). Xenobiotic metabolism has been extensively identified in different kinds of bacteria, which

are capable of degrading multiple kinds of xenobiotic compounds like aromatic compounds, phenolic compounds and dyes (Diaz et al., 2004).

1.3.3 Taxonomy

Early studies on bacterially mediated MC degradation are dominated by culture-dependent work (Cousins et al. 1996, Jones et al. 1994, Park et al., 2001). These studies have reported a number of MC-degrading bacterial isolates that are predominantly restricted within an alphaproteobacteria family, i.e. Sphingomonaceae (example genera are Sphingomonas,

Sphingopyxis and Sphingoscinicella) (Maruyuma et al., 2006; Jones et al., 1994; Hoefel et al.,

2009). An increasing number of studies have also shown to report the presence of MC degrading bacteria in betaproteobacteria and gammaproteobacteria (Chen et al., 2010; Krishnan et al., 2018;

Kumar et al., 2019). Probiotic bacteria like Lactobacillus rhamnosus strain GG and LC-705,

Bifidobacterium longum 46, Bifidobacterium lactis 420 and Bifidobacteium lactis Bb12 have also been shown to possess the ability to degrade MC in aqueous solutions (Nybom et al., 2007).

Mucilage of Microcystis cells have also been shown to possess the presence of MC degrading bacteria. The presence of these MC degrading bacteria present in mucilage are positively correlated to the concentration of MC (Maruyuma et al., 2003).

1.3.4 Factors impacting MC degradation

The rate of microbial degradation of MCs is affected by a number of abiotic factors including temperature, pH, DOC (Dissolved organic carbon) and nutrient availability (Phujomjai 7

et al., 2013; Manheim et al., 2018). Factors affecting the gene abundance of mlrA gene have also been studied under various nutrient conditions of nitrogen, phosphorous and peptone (Li et al.,

2011). Some of the growth conditions (pH, temperature, organic nutrient) for MC degrading bacteria (MC-degrading bacteria) are determined by the conditions seen in eutrophic lakes.

Aquatic monitoring has revealed that the pH and the temperature of the upper water layer in eutrophic water systems increases to 9-10 and 30-35ºC respectively (Lehman. 2007). This shift in the temperature and pH occurs during the sudden proliferation of water blooms because of cyanobacterial photosynthesis (Lopez-Archilla et al. 2004; Lehman 2007; EI Herry et al. 2008;

Okell et al., 2009). Thus, some MC degrading bacteria are likely to be active, or proliferate in an alkaline pH range and high temperature, to rapidly degrade MCs. This hypothesis is partly supported by the finding that MC is degraded better under alkaline conditions and increased temperature by mixed bacteria in water samples obtained during toxic water blooms (Saitou et al.,

2003). However, other MC-degrading bacteria, such as Novosphingobium sp. strain MD-1, have been found to only grow under conditions of neutral pH but not under alkaline conditions (Saitou et al., 2003). Studies have also found that temperatures that are lower or higher than favorable for optimum growth, bacterial metabolism and the production of biodegradation enzymes are less active, therefore slowing the rate of MC degradation (Somdee et al., 2013). Eutrophic lakes show excessive organic nutrient content or the elements carbon, nitrogen and phosphorus (Carmichael et al., 1992). Biodegradation of MCs in the presence of these excessive nutrients needs to be determined for future application of MC-degrading bacteria (Carmichael et al., 1992). Therefore, 8

abiotic factors of pH, temperature and the presence of organic sources of carbon or nitrogen have different impact on degradation of MCs by different bacterial taxa (Phujomajai et al., 2013).

Characterizing an MC-degrading bacterium that can grow under alkaline pH conditions (9-10) and high temperatures (30-35ºC) would give a better understanding on how microbes degrade MCs during the appearance of blooms. This would further contribute to a better understanding of MC dynamics in aquatic environments.

1.4 Toxicity of MCs

1.4.1: Mode of MC Exposure

MCs contamination in natural waters can reach human via various routes, including chronic and accidental ingestion of contaminated drinking water; inhalation or contact with the nasal mucous membrane, and dermal contact with toxins during recreational activities consumption of contaminated vegetables and fruits irrigated with water containing cyanotoxins; consumption of aquatic organisms (fish, shellfish, etc.) (Watanabe et al., 1985; Poste et al., 2011). Consumption of this toxin at concentration of 10 µg/L has been found to cause episodes of vomiting, diarrhea, abdominal pain and blistering around the mouth (US, EPA, 2013). The International Agency for

Research on Cancer classifies MC-LR as a potential carcinogen due to its ability to cause hepatic cancer (Falconer et al., 1992).

MCs (MCs) are rarely ingested directly in high lethal acute doses, but chronic frequent exposure to the toxin leads to illness in humans and animals alike (Backer et al., 2008). A number 9

of public health events have involved of MC contamination in the hospital use, drinking and recreational waters (Carmichael et al., 1992). Bioaccumulation of the toxins have been studied previously by many authors (Tencalla et al., 1994; Vasconcelos, 1995; Williams et al.,

1997; Amorim and Vasconcelos, 1999; Thostrup and Christoffersen, 1999). Crustaceans and fishes from Sepitaba Bay in Brazil have shown bioaccumulation of the toxin in lower concentrations, but enough to contaminate the aquatic animals (Magalhaes et al., 2003). Aquatic plants also show bioaccumulation of the toxin (Mitrovic et al., 2005) along with mussels showing the presence of the toxins in their muscles (Williams et al., 1997). Consuming these aquatic animals which bioaccumulate MCs is lethal and a number of studies have described the dangers it poses (Xie et al., 2005) like hepatocellular carcinoma and hepatic cell death.

1.4.2: Mechanism of toxicity

Once the MCs enter the cell they bind to OATP (organic anion transporter protein) and inhibit Protein phosphatases 1 and 2a (PP1 and 2a) (Steiner et al., 2016; Fischer et al., 2005).

Abundance of this receptor on the liver cells makes MCs in drinking water as a risk factor for primary liver cancer (PLC). Hepatic cancer incidence reported in China and Brazil has been related to MC contamination in drinking water resources (Jochimsen et al., 1998, Ueno et al., 1996).

Therefore, International Agency for Research on Cancer has classified MCs as possible human carcinogens due to their inhibition of protein phosphatases, which leads to the hyper-

phosphorylation of cellular protein (Lone et al., 2015; Chen et al., 2009). Since they cause hepatic cancer, MCs are also recognized as hepatotoxins or liver toxins (Scvircev et al., 2010).

Liver possesses the receptors OATP in abundance, required for the transport and binding of MC which makes it the most important target organ of MCs for animals (Fisher et al., 2005).

Lipid peroxidation, mitochondrial permeability transition (MPT), apoptosis and an increase in reactive oxidative species (ROS) are seen as a result of oxidative stress and measured by hydrogen peroxide release in MC-LR exposed hepatocytes when compared to control cells (Ding et al.,

2006). Apoptosis is the primary form of hepatocellular cytotoxicity observed as a result of MC- exposure.

1.4.3: Apoptotic pathway mechanism

Apoptosis is the form of programmed cell death observed as a result of bacterial or viral infection (Weng et al., 2007). Apoptosis can be induced by caspases independent or caspases dependent pathway. Caspases are cysteine proteases which play an indispensable role in triggering apoptosis (Decordier et al., 2005). Caspases-dependent pathway of apoptosis can either be extrinsic or intrinsic (Hongmei et al., 2012). Extrinsic pathway is induced by Fas and FasL, where

FasL belongs to the family of cytokines, and Fas is the death receptor present on the cell undergoing apoptosis (Wajant, 2002). FADD is also known as the Fas associated death domain, which is a complex formed after the association of Fas and FasL which in combination with caspases 8 gives DISC (Death inducing signaling complex), activating the downstream 11

procaspases for induction of cell death. (Ghobrial et al., 2005)(Fig 2). Intrinsic apoptotic pathway shows the presence of cytochrome c (cyt c) and Apoptotic protease activating factor-1 (Apaf-1) released by mitochondria which activates caspases 8 and the other downstream caspases for apoptosis induction (Scoltock et al., 2004; Fulda et al., 2004). The extrinsic and intrinsic pathways both end at the point of the execution phase, considered the final pathway of apoptosis (Salvesen,

2002). Execution caspases activate cytoplasmic endonuclease, which degrades nuclear material, and proteases that degrade the nuclear and cytoskeletal proteins (Walsh et al., 2008). It is the activation of the execution caspases 3, 6 and 7 that ultimately leads to cell death. Fas and Fas-L induced extrinsic apoptotic pathway has been the most commonly observed apoptotic pathway

(Ghobrial et al. 2005).

Apoptosis (intrinsic/extrinsic) is morphologically associated with nuclear chromatin condensation, cellular shrinkage, and cytoplasmic condensation, detachment of the cells from surrounding cells, membrane blebbing and the formation of apoptotic bodies (Ashkenazi and Dixit,

1999; Saraste, 1999). Apoptosis also leads to DNA fragmentation and release of proteases (Fiers et al., 1999; Higuchi et al., 2003). These morphological changes have been studied in vitro for

MC-LR induced apoptosis in hepatic cells (Mankiewicz et al., 2001; Falconer et al., 1992).

1.4.4: Current knowledge on MC induced cell death

In comparison with hepatic cells, non-hepatic cells require a greater MC-LR dose and longer exposure time for cell death to occur (McDermott et al., 1998). MC-LR induced cell death in both 12

hepatic and non-hepatic cells has been found to be triggered by caspase-3 (Zhang et al., 2010). But it is still not clear whether the MC-LR induced hepatotoxicity is based on reversible cytoskeletal derangement (Eriksson et al., 1989), secondary hepatocyte necrosis due to insufficient blood supply or apoptotic induced cell death. Therefore, the proposed work aims to understand the biological complexity of induction of apoptosis by investigating the expression of proteins multiplexed with morphological changes seen after administration of MC.

MC exposure induces the production of reactive oxidative species (ROS), which leads to mitochondrial permeability potential (MPP) and apoptosis via the intrinsic pathway (Weng et al.,

2007; Huang et al., 2016; Li et al., 2016). Studies have also suggested the presence of extrinsic pathway via the NF-kB pathway (Feng et al., 2011) in INS-1 cells (Ji et al., 2011). Due to the lack of clarity regarding the pathway adopted by MC-LR treated cells, further works needs to be performed to elucidate the cytotoxicty of MC.

Hypothesis and Objectives

Hypothesis 1: Lake Erie harbor a diversity of culturable MC-degrading bacteria.

Rationale: Lakes with a history of cyanobacterial blooms harbor a diversity of MC degrading bacteria. Therefore, Lake Erie being frequently exposed to toxic cyanobacterial blooms, is likely to harbor MC-degrading bacteria (Mou et al., 2013). A recent culture-independent metagenomic study conducted on the metagenomics identification of MC-degrading bacteria from Lake Erie 13

have found a heterogeneous population consisting of phyla Actinobacteria, Bacteroidetes,

Firmicutes, Planctomycetes, Proteobacteria of the alpha, beta, gamma and delta/epsilon subdivisions, and Verrucomicrobia (Mou et al., 2013) . Therefore, unlike the previous studies

(Manage et al., 2009), which identified most MC-degrading as Sphingomonas and related taxa, we expect to recover taxonomically diverse MC-degrading bacteria from Lake Erie.

Objective 1.1: To obtain pure culture isolates of MC-degrading bacteria from Lake Erie.

Objective 1.2: To physiologically and taxonomically identify MC-degrading bacteria.

Objective 1.3: To identify the effects of abiotic and biotic factors on MC-degradation.

Hypothesis 2: MC-degrading bacteria from Lake Erie possess an alternative pathway to break down MC-LR.

Rationale: A metagenomic analysis performed on bacteria from Lake Erie seemed to show lower abundance of the mlrA gene required for MC degradation (Mou et al., 2013). The study also proposed the presence of an alternative xenobiotic MC- degradative pathway in addition to the mlr based cleavage pathway (Mou et al., 2013). Therefore, this study will look into identifying this novel MC-degrading pathway in MC-degrading bacteria isolated from Lake Erie.

Objective 2.1: To perform mlrA target PCR amplification for MC-degrading bacteria

Objective 2.2: To perform random mutagenesis for MC-degrading /mlrA- bacteria and identify the

MC degradation involved genes.

Objective 2.3: To identify the intermediates and products of MC degradation using LC-MS 14

Hypothesis 3: MC-LR triggers apoptosis in human liver cells via a dualistic mechanism involving the extrinsic and intrinsic apoptotic pathway.

Rationale: It has been shown that Fas and Fas-L along with caspase 3 and ROS (reactive oxidative species) participate in the induction of apoptosis for MC-LR treated HepG2 cells (Feng et al., 2011).

However the caspase expressed in the apoptosis induced by MC-LR is yet to be identified, which is required for identification of the apoptotic pathway adopted in MC-LR treated hepatic cells. Hence, this project will be identifying the specific caspase involved in the apoptosis of MC-LR treated cells and also the time of induction for every caspase activated to identify the trigger for the apoptotic pathways. This would in extension enable us to identify the apoptotic pathway adopted by MC-LR treated HepG2 cells. A human hepatoma cell line, HepG2, are found to be extremely similar to human hepatocytes in terms of the functions and proteins expression. This makes them an ideal model to be studied for liver toxicity and metabolism (Knasmuller et al., 1998).

Objective 3.1: To confirm the induction of apoptosis by MC-LR in HepG2 cells using microscopy and Alamar Blue assay.

Objective 3.2: To identify the caspase expressed in MC-LR treated HepG2 cells

Objective 3.3: To identify potential time lag between expressions of different proteins induced in

MC-LR treated HepG2 cells

1.5 Proposed work

The current work fills the above mentioned knowledge gaps regarding biological degradation of MC, novel MC degradation pathway and the apoptotic effects of MC on hepatic cell line. The first step of the study involved identifying MC degrading bacteria from Lake Erie and the factors influencing the degradation of MC (Chapter 2 and 3). Subsequent work was performed for identifying the genes and intermediates involved in MC degradation performed by bacteria from

Lake Erie (Chapter 4). Hepatic cells treated with MC were used to identify the proteins expressed and the apoptosis induced using microscopy and flow cytometry (Chapter 5). The data collected from these studies will be instrumental in developing a better understanding of the toxin MC in terms of its biological degradation and hepatotoxic effects.

Table 1: Highest microcystin concentration recorded for various lakes

Lakes Highest MC concentration (µg/L) (June-July) (EPA., 2015) Lake Taihu 10.38 Lake Erie 100 Lake Poyang 9.97 Lake Chaohu 10 Upper Kalamath Lake 1-17 Lake Houston <0.2 Grand Lake St Mary’s 2000 Conesus Lake 5.07 Silver Lake 10.716

Table 2: Previously isolated MC degrading bacteria

Organisms Phylum References Arthrobacter sp. Alpha proteobacteria Manage et al., 2009; Lawton et al., 2011 Bacilllus sp. Firmicutes Hu et al., 2012; Kansole and Lin, 2016 Brevibacterium sp. Actinobacteria Manage et al., 2009; Lawton et al., 2011 Burkholderia sp. Alpha proteobacteria Lemes et al., 2008 Methylobacillus sp. Betaproteobacteria Hu et al., 2009 Morganella morganii Gammaproteobacteria Eleuterio and Batista., 2010 Orchrabactrum anthropi Proteobacteria Ramani et al., 2011 Paucibacter toxinivorans Proteobacteria Rapala et al., 2005

Poeteriochromonas sp. Chrysophyta Zhang et al., 2008 Pseudomonas aeruginosa Gammaproteobacteria Takenaka and Watanabe, 1997

Ralstonia solanacearum Proteobacteria Yan et al., 2004 Rhodococcus sp. Actinobacteria Manage et al., 2009; Lawton et al., 2011 Sphingomonas sp 7CY Proteobacteria Ishii et al., 2004 Sphingomonas sp. ACM- Proteobacteria Jones et al., 1994; Bourne et al., 3962 1996 Sphingomonas sp. B9 Proteobacteria Harada et al., 2004; Imanishi et al., 2005 Sphingomonas sp CBA4 Proteobacteria Valeri et al., 2006 Sphingomonas sp. MD-1 Proteobacteria Saito et al., 2003 Sphingomonas sp. MDB2 Proteobacteria Maruyama et al., 2006

Sphingomonas sp. MDB3 Proteobacteria Maruyuma et al., 2006

Sphingomonas isolate Proteobacteria Somdee et al., 2013 NV-3 Sphingomonas sp. Y2 Proteobacteria Park et al., 2001; Maruyuma et al., 2003 Sphingopyxic sp. LH21 Proteobacteria Ho et al., 2007 Sphingopyxis sp. USTB- Proteobacteria Zhang et al., 2010 05 18

L. Rhamnosus LC-705 Firmicutes Nyborn et al., 2012 B. longum 46 Proteobacteria Nyborn et al., 2012 Stenotrophomonas sp. Proteobacteria Chen et al., 2010 EMS

Fig 1: Cyanobacterial blooms as seen in western basin of Lake Erie in the Summer of 2016

Fig 2: Schematic representation of mlr and xenobiotic degradation pathway

Fig 3: Schematic representation of the extrinsic and intrinsic apoptotic pathway

References

1. Amorim, A and Vasconcelos, V (1999). Dynamics of MCs in the mussel Mytilus

galloprovincialis. Toxicon 37(7): 1041-1052.

2. Ashkenazi, A and Dixit, V (1999). Apoptosis control by death and decoy receptors.

Current Opinion in Cell Biology 11(2):255-260.

3. Berg, K., Carmichael, W., Skulber, O., Benestad, C and Underdall, B (1987).

Investigation of a toxic water-bloom of Microcystis aeruginosa (Cyanophyceae) in Lake

Akersvatn, Norway. Hydrobiologia 144:97-103.Bláha, L., Pavel, B and Blahoslav M

(2009). Toxins produced in cyanobacterial water blooms - Toxicity and

risks. Interdisciplinary Toxicology 2(2):36-41.

4. Carmichael, W (1992). The toxins of cyanobacteria. Scientific American 2(7):78-86.

5. Chen, J., Xie, P., Li, L and Xu, J (2009). First Identification of the Hepatotoxic MCs in

the Serum of a Chronically Exposed Human Population Together with Indication of

Hepatocellular Damage. Toxicological sciences 108(1):81-89.

6. Chen, J., Li, B., Wei, Z., Shao, H., Jing, D., Yan, F and Zhi Q (2010). Degradation of

MC-LR and RR by a Stenotrophomonas sp. strain EMS isolated from Lake Taihu,

China. International Journal of Molecular Science 11: 896-911.

7. Christoffersen, K., Lyck, S and Winding, A (2002). Microbial activity and bacterial

community structure during degradation of MCs. Aquatic Microbial Ecology 27(2):125-

136. 22

8. Dai, R., Wang, P., Jia, P., Zhang, Y., Chu, X and Wang, Y (2016). A review on

factors affecting MCs production by algae in aquatic environments. World Journal

of Microbiology and Biotechnology: 32-51.

9. Decordier, I (2005). Influence of Caspase Activity on Micronuclei Detection: A Possible

Role for Caspase-3 in Micronucleation. Mutagenesis 20(3): 173-79.

10. Diaz, E (2004). Bacterial degradation of aromatic pollutants: a paradigm of metabolic

versatility. International Microbiology (2004)7:173-180.

11. Dietrich, D and Hoeger, S (2005). Guidance values for MCs in water and cyanobacterial

supplement products (blue-green algal supplements): a reasonable or misguided

approach? Toxicology and Applied Pharmacology 203(3): 273-289.

12. Dittmann, E., Neilan, B., Erhard, M., Dohren, H and Borner, T (2003). Insertional

mutagenesis of a peptide synthetase gene that is responsible for hepatotoxin production

in the cyanobacterium Microcystis aeruginosa PCC 7806. Molecular Microbiology

26(4):779-787.

13. Ding, X., Li, X., Hong-Ying, D., Ik-Kyo, C., and Jin-Ae Lee (2006). Toxic Effects of

Microcystis Cell Extracts on the Reproductive System of Male Mice. Toxicon 48(8):

973-79.

14. United States Environmental Protection Agency (EPA) (2015). Health Effects Support

Document for the Cyanobacterial Toxin Microcystins. EPA- 820R15102: 1-92.

15. Eriksson, E., Paatero, I., Meriluoto, A., Codd, A., Kass, E., Nicotera, P and Orrenius S

(1989). Rapid microfilament reorganization induced in isolated rat hepatocytes by MC-

LR, a cyclic peptide toxin. Experimental Cell Research.185: 86–100.

16. Eleuterio, L and Batista, J (2010). Biodegradation studies and sequencing of MC-LR

degrading bacteria isolated from a drinking water biofilter and a fresh water lake.

Toxicon 55(8):1434-1442.

17. Falconer, I and Yeung, D (1992). Cytoskeletal changes in hepatocytes induced

by Microcystis toxins and their relation to hyperphosphorylation of cell proteins.

Chemico-Biological Interactions 81(1-2):181-196.

18. Feng, G., Musa, A., Ying, Li and Yansheng Bai (2011). NF-κB Mediates the Induction

of Fas Receptor and Fas Ligand by MC-LR in HepG2 Cells. Molecular and Cellular

Biochemistry 352.1(2): 209-19.

19. Fiers, W., Beyaert, R., Declercq, W and Vandenabeele, P (1999). More than one way to

die: apoptosis, necrosis and reactive oxygen damage. Oncogene 18:7719-7730.

20. Fischer, W., Altheimer, S., Cattori, V., Meier, P., Dietrich, D and Hagenbuch, B (2005).

Organic anion transporting polypeptides expressed in liver and brain mediate uptake of

MC. Toxicology and Applied Pharmacology 203(3):257-263.

21. Fulda, S (2004). Sensitization for Tumor Necrosis Factor-Related Apoptosis-Inducing

Ligand-Induced Apoptosis by the Chemopreventive Agent Resveratrol. Cancer

Research 64(1): 337-46. 24

22. Fontanillo, M and Kohn, M (2018). MCs: Synthesis and structure–activity relationship

studies toward PP1 and PP2A. Bioorganic and Medicinal Chemistry 26(6):1118-1126.

23. Gan, N., Xiao, Y., Zhu, L., Wu, Z., Liu, J., Hu, C and Song, L (2011). The role of MCs

in maintaining colonies of bloom‐ forming Microcystis spp. Environmental

Microbiology 14(3):730-742.

24. Ghobrial, I., Witzig, T and Adjei, A (2009). Targeting Apoptosis Pathways in Cancer

Therapy. A Cancer Journal for Clinicians 55(3): 178-94.

25. Ginn HP, Neilan BA (2010). NtcA from Microcystis aeruginosa PCC 7806 is

autoregulatory and binds to the MC promoter. Appl. Environ. Microbiol. 76:4362–4368.

26. Harada, K. Chemistry and detection of MCs in Toxic MCs, M. F. Watanabe, K.

I. Harada, W. W. Carmichael, and H. Fujiki, Eds., 103–143, CRC Press, Boca

Raton, Fla, USA, 1996.

27. Harada, K.-I., Imanishi, S., Kato, H., Masayoshi, M., Ito, E., Tsuji, K( 2004). Isolation

of Adda from MC-LR by microbial degradation. Toxicon 44:107-109.

28. Hesse, K., Kohl, J.-G. (2001) Effects of light and nutrient supply on growth and MC

content of different strains of Microcystis aeruginosa. p. 104-115.

29. Hisbergues, M., Christiansen, G., Rouhiainen, L., Sivonen, K and Borner T (2003).

PCR-based identification of MC-producing genotypes of different cyanobacteria

genera. Archives of Microbiology 180:402-410.

30. Ho, L., Gaudieux, A., Fanok, S., Newcombe, G and Humpage, A (2007). Bacterial

degradation of MC toxins in drinking water eliminates their toxicity. Toxicon 50(3):438-

441.

31. Ho, L., Sawade, E and Newcombe, G (2012). Biological treatment options for

cyanobacteria metabolite removal – A review. Water Research 46(2012): 1536-1548.

32. Hoefel, D., Adriansen, C., Bouyssou, M., Saint, C., Newcombe, G and Ho L (2009).

Development of an mlrA gene directed TaqMan PCR assay for quantitative assessment

of MC- degrading bacteria within water treatment plant sand filter biofilms. Appl

Environmental Microbiology 75:5167-5169.

33. Hongmei, Z (2012). Extrinsic and Intrinsic Apoptosis Signal Pathway Review. Intech:1-

12.

34. Hu, L., Yang, J., Zhou,W., Yin, Y., Chen, J and Shi, Z (2009). Isolation of

a Methylobacillus sp. that degrades MC toxins associated with cyanobacteria. New

Biotechnology 26(3-4):205-211.

35. Hu, L., Zhang, F., Li, C and Wang, M (2012). Biodegradation of MCs by Bacillus sp.

strain EMB. Energy Procedia 16:2054-2059.

36. Imanishi, S., Kato, H., Mizuno, M., Tsuji, K., Harada, K.-I.(2005). Bacterial degradation

of MCs and nodularin. Chemical Research in Toxicology 18:591-598.

37. Ishii, H., Nishijima, M., Abe, T (2004). Characterization of degradation process of

cyanobacterial hepatotoxins by a Gram-negative aerobic bacterium. Water Research

38:2667-2676.

38. Jahnichen S, Ihle T, Petzoldt T, Benndorf J (2007). Impact of inorganic carbon

availability on MC production by Microcystis aeruginosa PCC 7806. Applied and

Environmental Microbiology 73(21):6994-7002.

39. Jahnichen, S., Long, B and Petzoldt, T (2011). MC production by Microcystis

aeruginosa: Direct regulation by multiple environmental factors. Harmful Algae 12:95-

104.

40. Jannsen, D., Dinka, I., Poelarends, G and Terpstra, P (2005). Bacterial degradation of

xenobiotic compounds: evolution and distribution of novel enzyme activities.

Environmental Microbiology 7(12):1868-1882.

41. Jiang, Y., Shao, J., Wu, X., Xu, Y and Li, R (2011). Active and silent members in

the mlr gene cluster of a MC-degrading bacterium isolated from Lake Taihu, China.

FEMS Microbiology letters 322(2):108-114.

42. Jochimsen, E., Carmicheal, W., An, J and Cardo, D (1998). Liver Failure and Death

after Exposure to MCs at a Hemodialysis Center in Brazil. The New England Journal of

Medicine 338:873-878.

43. Jones, G and Orr, P (1994). Release and degradation of MC following algicide treatment

of a Microcystis aeruginosa bloom in a recreational lake, as determined by HPLC and

protein phosphatase inhibition assay. Water Research 28:871-876.

44. Li, Y., Shao, J., Wu, X., Xu, Y and Li, R (2011). MC production by Microcystis

aeruginosa: Direct regulation by multiple environmental factors. FEMS Microbiology

Letters 322(2):108-114.

45. Kansole, M and Lin, T (2016). MC-LR Biodegradation by Bacillus sp.: Reaction Rates

and Possible Genes Involved in the Degradation. Water 8(11): 508-520.

46. Kormas, K and Lymperopoulou, D (2013). Cyanobacterial toxin degrading bacteria:

who are they? BioMedical Research International 2013 (2013): 1-12.

47. Labine, M and Minuk, G (2009). Cyanobacterial toxins and liver disease. Canadian

Journal of Physiology and Pharmacology 87(10):773-788.

48. Lambert, T., Holmes, C and Hrudey, S (1994). MC class of toxins: health effects and

safety of drinking water supplies. Environmental Reviews 2(2): 167-186.

49. Lawton, L.A., Welgamage, A., Manage, P.M., Edwards, C., 2011. Novel bacterial

strains for the removal of MCs from drinking water. Water Science & Technology 63,

1137e1142.

50. Lehman, E (2007). Seasonal occurrence and toxicity of Microcystis in impoundments

of the Huron River, Michigan, USA. Water Research 41(4):795-802.

51. Lemes, G.A.F., Kersanach, R., Pinto, L.S., Dellagostin, O.A., Yunes, J.S., Matthiensen,

A(2008). Biodegradation of MCs by aquatic Burkholderia sp. from a South Brazilian

coastal lagoon. Ecotoxicology and Environmental Safety 69:358-365.

52. Lone, Y., Kohli, R and Bhide, M (2015). An overview of the toxic effect of potential

human carcinogen MC-LR on testis. Toxicology Reports 2(2015):289-296.

53. Lopez-Archilla, A., Moreira, D., Lopez-Garcia, P and Guerrero, C (2004).

Phytoplankton diversity and cyanobacterial dominance in a hypereutrophic shallow lake

with biologically produced alkaline pH. Extremophiles 8(2): 109-115.

54. Ma, G., Pei, H., Hu, W., Xu, X., Ma, C and Li, X (2014). The removal of cyanobacteria

and their metabolites through anoxic biodegradation in drinking water sludge.

Bioresource Technology 165(2014):191-198.

55. MacKintosh, C., Beattie, K., Klumpp, S., Cohen, P and Codd, G (1990). Cyanobacterial

MC‐ LR is a potent and specific inhibitor of protein phosphatases 1 and 2A from both

mammals and higher plants. FEBS Letters 264(2):187-192.

56. Magalhaes, V., Marinho, M., Domingo, P., Oilveira, A., Costa, S., Azevedo, L and

Azevedo, A (2003). MCs (cyanobacteria hepatotoxins) bioaccumulation in fish and

crustaceans from Sepetiba Bay (Brasil, RJ). Toxicon 42 (2003):289-295.

57. Manheim, D., Cheung, Y and Jing, S (2018). The Effect of Organic Carbon Addition on

the Community Structure and Kinetics of MC-Degrading Bacterial Consortia. Water 10

(1523): 1-22. 29

58. Manage, P., Edwards, C., Singh, B and Lawton, L (2009). Isolation and identification

of novel MC-degrading bacteria. Applied Environmental Microbiology 75:6924–6928.

59. Mankiewicz, J., Tarczynska, M., Fladmark, K., Doskeland, S., Walter, Z and Zaleswki,

M (2001). Apoptotic effect of cyanobacterial extract on rat hepatocytes and human

lymphocytes. Environmental Toixcology 16(3): 225-233.

60. Maruyama, T., Kato, K., Yokoyama, A., Tanaka, T., Hiraishi, A., Park, H.-D (2003).

Dynamics of MC-degrading bacteria in mucilage of Microcystis. Microbial Ecology

46:279-288.

61. Maruyama, T (2006). Sphingosinicella MCivorans gen. nov. sp. nov. a MC-degrading

bacterium. International Journal of Systematic and Evolutionary Microbiology 56(1):

85-89.

62. McDermott, C., Nho, C., Howard, W and Holton, B (1998). The cyanobacterial toxin,

MC-LR, can induce apoptosis in a variety of cell types. Toxicon 36(12):1981-1986.

63. Mitrovic, S., Allis, O., Furey, A and James, K (2005). Bioaccumulation and harmful

effects of MC-LR in the aquatic plants Lemna minor and Wolffia arrhiza and the

filamentous alga Chladophora fracta. Ecotxicology and Environmental safety

61(3):345-352.

64. Mou X., Lu, X., Jacob, J and Heath, R (2013). Metagenomic identification of

bacterioplankton taxa and pathways involved in MC degradation in Lake Erie. PLoS

ONE 8(4):1-14.

65. Nybom,S., Salminen, S and Meriluoto, J (2007). Removal of MC-LR by strains of

metabolically active probiotic bacteria. FEMS Microbiology Letters 270(1):27-33.

66. Peregrin-Alvarez, J., Sanford, C and Parkinson, J (2009). The conservation and

evolutionary modularity of metabolism. Genome Biology 10(63): 1-17.

67. Ramani, A., Rein, K., Shetty, K and Jayachandran, K (2012). Microbial degradation

of MC in Florida’s freshwaters. Biodegradation 23(1) 35-45.

68. Oh, H., Le, S., Jang, M.and Yoon, B (2000). MC Production by Microcystis aeruginosa

in a Phosphorus-Limited Chemostat. Applied and Environmental Microbiology

66(1):176-179.

69. Orr, P and Jones, G (2003). Relationship between MC production and cell division rates

in nitrogen‐ limited Microcystis aeruginosa cultures. Limnology and Oceanography

43(7): 1604-1614.

70. Paerl, H and Otten, T (2013). Harmful cyanobacterial blooms: causes, consequences,

and controls. Microbial Ecology 65(4): 995-1010.

71. Park, H., Sasaki, Y., Maruyama, T., Yanagisawa, E., Hiraishi, A., Kato, K (2001).

Degradation of the cyanobacterial hepatotoxin MC by a new bacterium isolated from a

hypertrophic lake. Environmental Toxicology 16(4): 337-343. 31

72. Pearson, L., Nihali, T., Moffitt, M., Kellmann, R and Neilan, R (2010). On the

chemistry, toxicology and genetics of the cyanobacterial toxins, MC, nodularin,

saxitoxin and cylindrospermopsin. Marine Drugs 8(5): 1650-1680.

73. Phujomjai, Y., Somdee, A and Somdee T (2015). Biodegradation of MC [Dha7] MC-

LR by a novel MC-degrading bacterium in an internal airlift loop bioreactor. Water

Science and Technology73:267-274.

74. Poste, A., Hecky, R and Stephanie J. Guildford (2011). Evaluating MC Exposure Risk

through Fish Consumption. Environmental Science & Technology 45 (13): 5806-5811.

75. Rapala, J., Berg, K.A., Lyra, C., Niemi, R.M., Manz, W., Suomalainen, S., Paulin, L.,

Lahti, K., 2005. Paucibacter toxinivorans gen. nov., sp. nov., a bacterium that degrades

cyclic cyanobacterial hepatotoxins MCs and nodularin. International Journal of

Systematic and Evolutionary Microbiology 55, 1563-1568.

76. Rinehart, K., Harada, K., Namikoshi, M., Chen, C., Harvis, C., Munro, M., blunt, J.,

Muligan, P and Beasley, V (1988). Nodularin, MC, and the configuration of Adda.

Journal of the American Chemical Society 110(25):8557-8558.

77. Rinta-Kanto, J and Wilhelm, S (2006). Diversity of MC-Producing Cyanobacteria in

Spatially Isolated Regions of Lake Erie. Applied and Environmental Microbiology

72(7):5083-5085.

78. Rinta-Kanto, J., Konopko, E., DeBruyn, J., Bourbonniere, R., Boyer, G and Wilhelm, S

(2009). Lake Erie Microcystis: Relationship between MC production, dynamics of

genotypes and environmental parameters in a large lake. Harmful Algae 8(5):665-673.

79. Saito, T., Okano, K., Park, D., Itayama, T., Inamori, Y., Neilan, A., Burns, P and Suquira

N (2003). Detection and sequencing of the MC LR- degrading gene, mlrA from new

bacteria isolated from Japanese lakes. FEMS Microbiology Letters 229:271-276.

80. Salvesen, G (2002). Caspases and Apoptosis. Essays in Biochemistry 38:9-19.

81. Saraste, A (1999). Morphologic criteria and detection of apoptosis. Herz 24(3):189-195.

82. Schmidt, J., Wilheml, S and Boyer, G (2014). The fate of MCs in the environment and

the challenges of monitoring. Toxins 6(12): 3354-3387.

83. Scoltock,A and Cidlowski, J (2004). Activation of intrinsic and extrinsic pathways in

apoptotic signaling during UV-C-induced death of Jurkat cells: the role of caspase

inhibition. Experimental Cell Research 297 (1): 212-223.

84. Svircev, Z., Baltic, V., Gantar, M., Jukovic, M., Stojanovic, D and Baltic, M (2010).

Molecular aspects of MC-induced hepatotoxicity and hepatocarcinogenesis. Journal of

Environmental Science and Health Part C Environmental Carcinogenesis &

Ecotoxicology Reviews 28(1):39-59.

85. Sedmak, B and Kosi, G (1998). The role of MCs in heavy cyanobacterial bloom

formation. Journal of Plankton Research 20(4):691-708.

86. Shimizu, K., Maseda, H., Okano, K., Hiratsuka, T., Jimbo, Y., Xue, Q., Akasako, H.,

Itayama, T., Utusumi, M., Zhang, Z and Suguira, N (2013). Determination of MC-LR

degrading gene mlrA in biofilms at a biological drinking water treatment facility. Maejo

International Journal of Science and Technology 7(special issue):22-35.

87. Sigee, R., Glenn, M., Andrews, E., Bellinger, R., Butler, H., Epton, R and Hendry

(1999). Biological control of cyanobacteria: principle and possibilities. Hydrobiologia

395(396):161-172.

88. Somdee, T., Thunders, M., Rucks, M., Allison, M and Page R (2013) Degradation of

[Dha (7)] MC-LR by a MC degrading bacterium isolated from Lake Rotoiti, New

Zealand. ISRN Microbiology 2013:1-8.

89. Springer-Verlag. Berlin.Higuchi, Y (2003). Chromosomal DNA fragmentation in

apoptosis and necrosis induced by oxidative stress. Biochemical Pharmacology

66(8):1527-1535.

90. Steiner, K., Zimmermann, L., Hagenbuch, B and Dietrich, D (2016). Zebrafish-Oatp

mediated transport of MC-congeners. Archives of Toxicology 90(5): 1129-1139.

91. Tencalla, F., Dietrich, D and Christian, S (1994). Toxicity of Microcystis aeruginosa

peptide toxin to yearling rainbow trout (Oncorhynchus mykiss). Aquatic Toxicology

30(3): 215-224.

92. Thostrup, L and Kirsten, C (1999). Accumulation of MC in Daphnia magna feeding on

toxic Microcystis. Archiv fur Hydrobiologi 145(4):447-467. 34

93. Tillett D, Dittmann E, Erhard M, von Döhren H, Börner T, Neilan BA (2000). Structural

organization of MC biosynthesis in Microcystis aeruginosa PCC7806: an integrated

peptide–polyketidesynthetase system. Chemical Biology 7:753–764.

94. Tonk, L (2005). The MC composition of the cyanobacterium Planktothrix agardhii

changes toward a more toxic variant with increasing light intensity. Applied

Environmental Microbiology 71:5177–5181.

95. Tsuji, K., Naito, S., Kondo, F., Ishikawa, N., Watanabe, M., Suzuki, M and Harada,

K (1994). Stability of MCs from cyanobacteria: Effect of light on decomposition and

isomerization. Environmental Science and Technology 28 (1994):173-177.

96. Van der Westhuizen, A and Eloff, J (1985). Effect of temperature and light on the

toxicity and growth of the blue-green alga Microcystis aeruginosa (UV-006). Planta

163(1):555-59.

97. Valeria, A.M., Ricardo, E.J., Stephan, P., Alberto, W.D (2006). Degradation of MC-RR

by Sphingomonas sp. CBA4 isolated from San Roque reservoir (Co´ rdoba - Argentina).

Biodegradation 17:447-455.

98. Wajant, H (2002). The Fas Signaling Pathway: More Than a Paradigm. Scienc

296(5573): 1635-1636.

99. Walsh, G., Culen, S., Sheridan, C., Luthi, A., Gerner, C and Martin, S (2008).

Executioner caspase-3 and caspase-7 are functionally distinct proteases. Proceedings of

the National Academy of Sciences of the United States of America 105(35):12815-

12819.

100. Watanabe, M and Oishi, S (1985). Effects of Environmental Factors on Toxicity of

a Cyanobacterium (Microcystis aeruginosa) under Culture Conditions. Applied and

Environmental Microbiology 49(5):1342-1344.

101. Weng, D., Yan, L., Yinna, W., Ying, L and Pingping, S (2007). The Role of ROS

in MC-LR-induced Hepatocyte Apoptosis and Liver Injury in Mice. Toxicology 232(1-

2): 15-23.

102. Wicks, R and Thiel, P (1990). Environmental factors affecting the production of

peptide toxins in floating scums of the cyanobacterium Microcystis aeruginosa in a

hypertrophic African reservoir Environmental Science & Technology, 24 (1990),

pp. 1413-1418.

103. Williams, D., Dawe, S., Kent, M., Andersen, R., Craig, M and Holmes, C (1997).

Bioaccumulation and clearance of MCs from salt water mussels, Mytilus edulis, and in

vivo evidence for covalently bound MCs in mussel tissues. Toxicon: Official Journal of

the International Society on Toxicology 35(11):1617-1625. S

104. Yang, F., Zhou, Y., Yin, L., Zhu, G., Liang, G and Pu, Y (2014). MC-Degrading

Activity of an Indigenous Bacterial Strain Stenotrophomonas acidaminiphila MC-LTH2

Isolated from Lake Taihu. PLOS One 9(1): 1-7.

105. Zhang, G., Pan, and Yan, H (2010). Microbial biodegradation of MC-RR by

bacterium Sphingopyxis sp. USTB-05. Journal of Environmental Sciences 22 (2):168-

175.

Chapter II: Isolation and characterization of microcystin degrading bacteria from Lake Erie

Preface:

This project was published under the title “Isolation and Characterization of

Microcystin-Degrading Bacteria from Lake Erie” in Bulletin of Environmental Contamination and Toxicology” Volume 101:1-7. My co-author, Yu- Quin Zhang helped us obtain pure culture isolates from Lake Erie and devise a protocol for isolation of pure culture isolates from Lake Erie water and sediment samples. The second co-author for my paper Xiaozhen Mou, who is also my dissertation advisor, helped me immensely with the design of the experiments, execution and writing of the paper.

Abstract

Heterotrophic bacteria are suggested as the major agents that degrade microcystins (MCs), a major cyanotoxins, in natural environments. However, little is known of the taxonomic and functional diversity of MC-degrading bacteria in Lake Erie of the Laurentian Great Lakes, the largest freshwater system on earth. This study obtained numerous bacterial cultures from Lake Erie and six of these showed the ability for using MCs. MC degradation rate of the isolates were impacted by temperature and pH. The key gene for MC degradation (mlrA) were absent from for all 6 MC degraders, indicating a novel degradation pathway. In addition for potentials used in MC bioremediation, two isolates may offer extra benefits as biofertilizers.

Introduction

Cyanobacterial harmful blooms (CyanoHABs) frequently develop in eutrophic waters and lead to multiple negative effects (Paerl et al., 2013). One of the harmfulness is the production of cyanotoxins, which cause significant ecological and health concerns, especially in waters that serve as drinking water supplies and recreational grounds (Blaha et al. 2009; Paerl et al. 2013).

Cyanotoxins produced during freshwater CyanoHABs are dominated by a group of liver toxins known as microcystins (MCs; Hisbergues et al. 2003). Over 90 MC isoforms have been identified; microcystin-leucine arginine (MC-LR) is the most abundant and widely studied one (Paerl et al.,

2013). MC-LR and all other MC isoforms possess a monocyclic heptapeptide structure, which makes them highly resistant to physical and chemical breakdowns in natural environments (Chen et al. 2010). MC degradation in nature is primarily carried out by heterotrophic bacteria (Kormas et al. 2013).

Early studies on MC biodegradation are dominated by culture-dependent work have reported a number of MC-degrading bacterial isolates that are predominantly restricted within the

Sphingomonaceae family (such as Sphingomonas, Sphingopyxis and Sphingoscinicella) (Jones et al. 1994; Maruyuma et al. 2006; Hoefel et al. 2009). Using Sphingomonas sp. ACM-3962 as a model system, MC-LR degradation was first identified to follow a step-wise cleavage process that is initiated by microcystinase (MlrA)(Bourne et al. 1996), therefore, its gene (mlrA) has been widely adopted to probe MC-degrading bacteria in various natural lakes (Somdee et al. 2013) and 39

man-made environments (Hoefel et al. 2009). However, the ubiquity of Sphingomonaceae and mlr genes to MC degradation is in question. Recent studies have identified many non-

Sphingomonaceae MC degraders, and many of them do not carry mlrA genes (Manage et al. 2009;

Somdee et al. 2013; Yang et al. 2014). Moreover, a recent metagenomic study indicates that a diverse group of bacterial taxa, especially those affiliated with betaproteobacteria, and xenobiotic metabolism-related genes may govern MC degradation in Lake Erie (Mou et al. 2013b). However, no direct evidence is available.

Lake Erie is the shallowest and most southern lake among the five Laurentian Great Lakes, which represent the largest surface freshwater system on earth. However, like most surface freshwaters, Lake Erie has become eutrophic and host periodic blooms of MC-producing

CyanoHABs (Michalak et al. 2013). A bloom event in Lake Erie in August 2014 resulted in a shutdown of public water supply from 500,000 residents (Henry 2014). Due to the obvious ecological, economic and health significance, MC transformation has been a hot research and remediation topic in Lake Erie and CyanoHAB-impacted freshwaters worldwide. However, majority studies have been focused on mechanisms that regulate the synthesis of MCs, leaving MC degradation relatively understudied.

This study aimed to obtain one of the first collections of MC-degrading bacterial pure cultures from Lake Erie and examine their potential diversity in taxonomy, phenotype, physiology and MC-degrading efficiency and pathway (i.e., whether or not involve mlr genes).

Materials and Methods

Bacterial isolation

Samplings were performed once every month in Lake Erie from July to September 2013 in areas with frequent and long-lasting CyanoHABs (HAB Bulletin, NOAA, 2017). Surface water samples were collected and filtered through 5 μm pore-size membrane filters (Whatman membrane filters, Pittsburgh, PA, USA). The resulting filtrates for each sample were amended with NH4NO3 and KH2PO4 (both at 0.05 mM, final concentrations) and incubated in the dark at 30 °C with a rotation rate of 200 rpm for 7 days. Afterwards, the incubated water was amended with 1 μg/ml

MC-LR (Cayman Chemicals, Ann Arbor, Michigan) and incubated at the same condition for another 7 days. Aliquots of 50 μl of incubated water were then spread onto agar plates prepared with R2A, IPS-I or LB media and incubated at 30°C in the dark for 48 hrs. Single colonies were re-streaked for at least five times to obtain pure isolates.

Screening for MC-degrading activities

Bacterial pure cultures were re-inoculated in liquid R2A media and incubated at the same condition for 48 hours. Cells were harvested by centrifugation and washed three times with PBS before being re-suspended in BG11 media and incubated without carbon source for 24 hrs. The starved cells were transferred into BIO-LOG MT2 plates (Hayward, CA, USA) containing MC-

LR (0, 0.1, 1 and 10 µg/ml; final concentrations). Samples without bacterial cells or MC-LR were also included. The MT2 plates were incubated in the dark at 30°C for a total of 48 hrs. Color 41

changes were tracked by measuring the absorbance at 630 nm using a Synergy Multimode Reader

(Biotech, VT, USA).

MC-LR degradation was also confirmed by growing carbon-starved bacterial pure cultures in 1 µg/ml of MC-LR in BG11 media. Bacterial growth was tracked by measuring optical density

(OD600) during the incubation. MC-LR consumption in the samples was also measured using a

Microcystins Nodularins ADDA ELISA kit (ABRAXIS Warminster, PA, USA) following the manufacturer’s protocol.

Effect of temperature and pH on degradation.

Carbon-starved cells were prepared following procedure described above and mixed with

BG11 media that contain 5 µg/ml (final concentration) of MC-LR. Cells were then incubated at various temperatures or pH and tracked for MC degradation as above. pH for the different suspensions were measured using a pH meter (pH 123, Hannah Instruments, MI, USA). If the pH was higher than desired, it was adjusted using sodium hydroxide solution, if the pH was lower than desired, it was adjusted using hydrochloric acid solution.

Phenotypic characterization of isolates.

Phenotypic characteristics of putative MC-degradation bacterial isolates, including colony pigmentation, cell morphology and gram nature, were examined using standard procedures (Clark et al. 1976). Motility of strains was examined by wet mount and Leifson’s staining technique for observation of the presence of flagella and cell movement using light microscopy (Clark et al.

1976). 42

16S rRNA and mlrA gene analysis.

Genomic DNAs of pure isolates were extracted using an UltraClean GelSpin DNA

Extraction Kit (MoBio, CarlsBad, CA, USA). 16S rRNA genes were amplified with primers 27F

(5´-AGAGTTTGATCCTGGCTCAG-3´) and 1492R (5´-TACGGYTACCTTGTTACGACTT-3´) following a PCR program that included 95 ºC for 3 mins, followed by 30 cycles at 95 ºC for 1 min, at 59 ºC for 1 min, at 72 ºC for 1 min, and a final step at 72ºC for 10 mins. PCR amplicons were examined using agarose gel (1%) electrophoresis and then purified using an Ultra Clean Gel

Purification Kit (MoBio, Carlsbad, CA, USA) before sequenced for near-full length at the

Macrogen Corporation, USA.

PCR amplification of mlrA gene were used the primers MF (5´-

CCTCGATGACCTCGTAGC-3´) and MR (5´-CGGCCATCTTCAGCAAT-3´ ) following a PCR program that included 94°C for 1 min followed by 30 cycles at 94°C for 20s, at 60°C for 10s, and at 72°C for 30 s (Saito et al. 2003). PCR amplicons were examined using agarose gel (1%) electrophoresis. A synthetic positive control of mlrA-containing plasmid was obtained from the

USGS Michigan-Ohio Water Science Center (Dr. D. France).

Statistical analysis.

All statistical analyses were performed using R (R core team, 2013). T test was used to compare the absorbance values among the different MC-LR treatments in the MT2 BIOLOG assay. Two-way Analysis of Variance (ANOVA) was used to compare bacterial growth and MC-

LR consumption during the growth assays under different temperatures and pH values. Pearson’s 43

product moment correlation analysis was performed to examine potential correlations between the

MC-LR degradation rate and temperature or pH.

Nucleotide sequence accession numbers.

Sequences of near-full-length (>1300 bp) 16S rRNA genes for six MC-degrading isolates were obtained from Macrogen corporation, USA and deposited into the NCBI GenBank under the accession numbers of KX185385-KX185387, KX185392, KX185397 and KX753361.

Results and Discussion

Isolation and screening for microcystin-degrading bacteria.

The culturing effort stopped when obtained a total of 500 bacterial strains from Lake Erie water samples. Six of these isolates showed increased absorbance values in BIOLOG MT2 assays when supplied with MC-LR (10 µg/ml) as the single carbon source (P < 0.05; Fig. 4). Meanwhile, corresponding negative controls of these six isolates maintained low absorbance values (Fig. 4).

These six bacteria were putatively identified as MC-degrading bacteria. When MC-LR supply was reduced to 1 µg/ml, the degradation rate of all six isolates significantly reduced and at 0.1

µg/ml, only half of the isolates still showed positive consumption of MCs within 48 hrs. (P < 0.05).

This indicate that substrate availability affects MC-degradation rate of bacteria and that bacteria may have varied affinity to MC-LR.

The MC-degradation ability of the six putative MC-degrading strains were confirmed by growth assays. With the same starting OD600 value (0.5) and volume (5 ml) and incubation conditions, all six bacterial isolates fully consumed MC-LR (Fig. 5). Coincident with consumptions of MC-LR, bacterial cell number significantly increased for all of the tested isolates

(t test; P < 0.05). LEw-1278 showed a close to linear degradation of MC-LR over time, while the other five isolates had a faster consumption of MC-LR in the first 24 hrs. than the second 24 hrs.

Based on measurements at 24 hrs., the average MC-LR consumption rate for the six isolates was

0.03 µg/ml/hr.

Phenotypic and genotypic characterization of microcystin degrading bacteria.

Obtained MC-degrading bacterial isolates showed a range of phenotypic traits in cell colony pigmentation, cell morphology, and gram nature (Table 3).

In accordance with the diversity of phenotypic traits, near full-length 16S rRNA genes sequences revealed that the six MC-degrading bacterial isolates were broadly affiliated with different bacterial taxa (Fig. 6). Four isolates were affiliated proteobacteria, either in the beta

(LEW-2) or the gamma-classes (LEw-1033, LEw-1278 and LEw-2166). Isolate LEw-2 was affiliated with Acidovorax facilis of Burkholderiales (betaproteobacteria). Members of

Burkholderiales accounted for ~20% of bacterioplankton community in the western basin of Lake

Erie (Mou et al. 2013a) and have been suggested as important MC-degraders in Lake Erie (Mou et al. 2013b). This is the first report for a Acidovorax member to degrade microcystins, although several Acidovorax species have been reported to degrade complex cyclic organic compounds, 45

such as monocyclic and polycyclic aromatic hydrocarbons (PAHs) (Singleton et al. 2001). The species A. facilis is a common soil taxon and some strains are used as plant growth promoting

(PGP) inoculants in bio-fertilizers (such as Accomplish LM by Loveland Products) to increase the yield of corn and soybean crops (Adesmoye et al. 2017). This is partly because A. facilis carry nitrilases that can degrade organic nitrogen to carboxylic acid and ammonia, which in turn can be readily used by plants (Wu et al. 2007). Application of A. facilis-containing bio-fertilizers has been studied in cornfields of Northwestern Ohio (Lentz et al. 2015); runoff from these and other agriculture fields is a potential source of this taxon in Lake Erie.

For MC-degrading isolates that were affiliated with gammaproteobacteria, LEw-1033 and

LEw-2166 were both closely affiliated with Pseudomonas putida. Pseudomonas species, have shown abilities to degrade recalcitrant aromatic compounds (Guerin et al. 1995; Morono et al.

2004), including MC-LR (Yang et al. 2014). Consistent with our findings, mlrA has not been identified in any of the previously isolated MC-degrading gamma-proteobacteria species. Isolate

LEw-1278 was affiliated with Stenotrophomonas maltophila. A MC-degrading S. maltophila has also been isolated from Lake Taihu, China (Yang et al. 2014) with a slightly slower MC degradation rate.

We also obtained two Gram-positive MC-degrading isolates in the phylum of Firmicutes, i.e., LEw-1238 (Brevibacillus brevis) and LEw-2010 (Bacillus thuringenesis). These species can degrade multiple refractory compounds, such as polyethylene and pyrene (Alhassani et al. 2007;

Nehra et al. 2016), but this is the first time to show their MC degradation abilities. Like A. facilis 46

(LEw-2), B. brevis strains have been recommended to serve as PGP inoculants (Nehra et al. 2016).

Therefore, applying A. facilis and B. brevis strains to farmland may also offer extra benefits in bio- remediating microcystin contaminations, which is a raising concern in areas that use CyanoHAB impacted water for irrigation.

It is noted that none of the obtained isolates was affiliated with or close relative to the

Sphingomonaceae of Alphaproteobacteria (Fig. 6), which family contains most of the reported cultured MC-degrading isolates and perform MC degradation via mlr genes-based cleavage pathway (Ho et al. 2007). In our subsequent PCR analysis, mlrA genes were not detected in any of our six isolates. In contrast, PCR was consistently successful when amplifying mlrA from the positive control (data not shown). Therefore, both of the taxonomic and functional gene make up of our MC-degrading isolates suggest alternative pathway(s) exist and sometime even be more important than mlr-based cleavage in MC degradation in freshwater systems. This is consistent with the increasing evidence from both culture-dependent and independent studies (Mou et al.

2013b; Kansole et al. 2016). Our isolates can serve as models to elucidate the proposed novel pathways and genes involved in the breaking down of MC-LR.

Effects of temperature and pH on MC-LR degradation.

For each isolate, both of the bacterial growth activity and MC-LR degradation rate were positively correlated (r=0.75) with temperature (P < 0.05; Fig. 7). At 15 ˚C, only LEw-1238 showed significant MC-LR degradation. As incubation temperature increased (25ºC, 30˚C and

35˚C), isolates gradually increased MC-LR degradation rate and consumed 100% of added MC- 47

LR (5 µg/ml) in 48 hrs. MC-LR degradation activities were observed at the highest rate when pH values were at 7 and 8. The degradation activity was significantly inhibited at either higher or lower pH conditions (ANOVA; P <0.05), except for LEw-2. For LEw-2, MC-LR degradation rate was maintained at all tested basic pH conditions; MC degradation was only inhibited at acidic pH conditions.

The revealed effects of T and pH on MC degradation were consistent with findings of previous culture studies and field observations (Phujomjai et al. 2015; Zhang et al. 2015). Faster degradation rate at higher T cannot potentially offset the increased production of MC by cyanobacteria during warm weather conditions. The optimal pH for our Lake Erie isolates in MC-

LR degradation was at pH 7-8, which matches the typical pH range during CyanoHAB blooms in

Lake Erie (Mou et al. 2013a). In addition, since pH is typically maintained at a low basic range during drinking water treatment (Stevik et al. 1999), our isolates bacteria have the potential to serve as inoculant during the filtration step of water treatment to augment MC degradation.

Acknowledgements

We would like to thank Dr. J. Ortiz for assistance in collecting samples, Serghei

Lordachescu for helping with culturing work, and for helping measurements of pH and temperature. This work was supported by Ohio Department of Higher Education Harmful Algal

Bloom Research Initiative (R/HHT-4), Lake Erie Protection Fund (SG450-13, SG514-2017) and

Kent State University Research Council. 48

. Isolate ID Colony Shape Gram Flagella mlrA

Color nature (+/-)

LEw-2 Pink Cocci ˗ ˗ -

LEw-1033 Yellow Coccobacilli ˗ ˗ -

LEw-1238 White Cocci + + -

LEw-1278 Orange Bacilli ˗ ˗ -

LEw-2010 White Cocci + ˗ -

LEw-2166 White Coccobacilli ˗ ˗ -

Table 3.Gentotypic and Phenotypic characteristics of microcystin-degrading bacterial isolates obtained from water samples of Lake Erie

Fig 4: Changes of MC-LR concentration (left axis, solid gray line) and optical density OD600 (right axis, dash black line) in BG11 growth media supplied with 1 µg/ml of MC-LR and bacterial isolates.

Fig.5: A neighbor-joining phylogenetic tree based on partial sequence of 16S rRNA gene showing the relationship sequences of isolated MC-degrading bacteria and their closely related relatives. Gray circles are to label previously identified MC-degrading bacteria. The GenBank accession numbers of the sequences are shown in parentheses. Bootstrap values that are higher than 50% are show at the branch nodes (1,000 resampling). The scale bar represents 0.02 nucleotide substitutions per position.

Fig.6: Effect of temperature on MC-LR degradation by MC- degrading bacteria.

Fig .7: Effects of pH on degradation of MC-LR by MC-degrading bacteria.

References 1. Adesemoye A, Yuen G, Watts D (2017). Microbial Inoculants for Optimized Plant Nutrient

Use in Integrated Pest and Input Management Systems. In Probiotics and Plant Health.

Springer, Singapore.

2. Alhassani H, Ashraf S (2007). Efficient microbial degradation of Toluidine blue dye by

Brevibacillus sp. Dyes and Pigments 75:395-400.

3. Bláha L, Pavel B, Blahoslav M (2009). Toxins produced in cyanobacterial water blooms -

Toxicity and risks. Interdisciplinary Toxicology 2:36-41.

4. Bourne D, Jones G, Blakeley R, Jones A, Negri A, Riddles P (1996). Enzymatic pathway

for the bacterial degradation of the cyanobacterial cyclic peptide toxin microcystin-LR.

Applied Environmental Microbiology 62:4086–4094.

5. Chen J, Li B, Wei Z, Shao H, Jing D, Yan F, Zhi Q (2010). Degradation of microcystin-

LR and RR by a Stenotrophomonas sp. strain EMS isolated from Lake Taihu,

China. International Journal of Molecular Science 11: 896-911.

6. Clark W (1976). A simplified Leifson flagella stain. Journal of Clinical Microbiology 3:

632-634.

7. Guerin W, Boyd S (1995). Maintenance and induction of naphthalene degradation activity

in Pseudomonas putida and an Alcaligenes sp. under different culture conditions. Applied

Environmental Microbiology 61:4061-4068.

8. Henry T (2014). Water crisis grips hundreds of thousands in Toledo area, state of

emergency declared. Toledo Blade.

9. Hisbergues M, Christiansen G, Rouhiainen L, Sivonen K, Borner T (2003). PCR-based

identification of microcystin-producing genotypes of different cyanobacteria genera.

Archives of Microbiology 180:402-410.

10. Hoefel D, Adriansen C, Bouyssou M, Saint C, Newcombe G, Ho L (2009). Development

of an mlrA gene directed TaqMan PCR assay for quantitative assessment of microcystin-

degrading bacteria within water treatment plant sand filter biofilms. Applied

Environmental Microbiology 75:5167-5169.

11. Ho L, Hoefel D, Saint C, Newcombe G (2007). Isolation and identification of a novel

microcystin-degrading bacterium from a biological sand filter. Water Research 41:4685-

4695.

12. Jones G, Orr P (1994). Release and degradation of microcystin following algicide treatment

of a Microcystis aeruginosa bloom in a recreational lake, as determined by HPLC and

protein phosphatase inhibition assay. Water Research 28:871-876.

13. Kansole M, Lin T (2016). Microcystin-LR biodegradation by Bacillus sp. : Reaction rates

and possible genes involved in the degradation. Water 50:1-19.

14. Kormas K, Lymperopoulou D (2013). Cyanobacterial toxin degrading bacteria: who are

they? BioMedical Research International 2013: 1-12.

15. Lentz E (2015). Corn response to Accomplish LM in Northwestern Ohio. 2015 American

Society of Agronomy Annual Meeting. Minneapolis, MN, Nov 15-18. #130-12.

16. Manage P, Edwards C, Singh B, Lawton L (2009). Isolation and identification of novel

microcystin-degrading bacteria. Applied Environmental Microbiology 75:6924–6928.

17. Maruyama T (2006). Sphingosinicella microcystinivorans gen. nov. sp. nov. a microcystin-

degrading bacterium. International Journal of Systems and Evolutionary Microbiology 56:

85-89.

18. Michalak A, Anderson E, Beletsky D, Boland S, Bosch N, Bridgeman T and et al. (2013).

Record-setting algal bloom in Lake Erie caused by agricultural and meteorological trends

consistent with expected future conditions. Proc Natl Acad Sci U S A 110: 6448-6452.

19. Morono Y, Unno H, Tanji Y, Hori K (2004). Addition of aromatic substrates restores

trichloroethylene degradation activity in Pseudomonas putida F1. Applied Environmental

Microbiology 70: 2830-2835.

20. Mou X, Jacob J, Lu X, Robbins S, Sun S, Ortiz, J (2013a). Diversity and distribution of

free-living and particle-associated bacterioplankton in Sandusky Bay and adjacent waters

of Lake Erie Western Basin. Journal of Great Lakes Research, 39: 352-357.

21. Mou X, Lu X, Jacob J, Heath R (2013b). Metagenomic identification of bacterioplankton

taxa and pathways involved in microcystin degradation in Lake Erie. PLoS ONE 8:1-

14.e61890.

22. Nehra V, Saharan B, Choudhary M (2016). Evaluation of Brevibacilllus brevis as a plant

growth promoting rhizobacteria for cotton (Gossypium hirsutum) crop. Springer Plus 5:

2584-2588.

23. Paerl H, Otten T (2013). Harmful cyanobacterial blooms: causes, consequences, and

controls. Microbial Ecology 65: 995-1010.

24. Phujomjai Y, Somdee A, Somdee T (2015). Biodegradation of microcystin [Dha7] MC-

LR by a novel microcystin-degrading bacterium in an internal airlift loop bioreactor. Water

Science and Technology 73:267-274.

25. Saito T, Okano K, Park D, Itayama T, Inamori Y, Neilan A, Burns P, Suquira N (2003).

Detection and sequencing of the microcystin LR- degrading gene, mlrA from new bacteria

isolated from Japanese lakes. FEMS Microbiology Letters 229:271-276.

26. Singleton D, Furlong M, Rathbun S, Whitman W (2001). Quantitative comparisons of 16S

rRNA gene sequence libraries from environmental samples. Applied Environmental

Microbiology 67: 4374–4376.

27. Somdee T, Thunders M, Rucks M, Allison M, Page R (2013). Degradation of [Dha (7)]

MC-LR by a microcystin degrading bacterium isolated from Lake Rotoiti, New Zealand.

ISRN Microbiology 2013:1-8.

28. Stevik T, Ausland G, Hansen J, Jenssen P (1999). The influence of physical and chemical

factors on the transport of E. coli through biological filters for wastewater purification.

Water Research 33: 3701-3706. 57

29. Wu S, Fogiel A, Petrillo K, Hann E, Mersinger L, DiCosimo R, O'Keefe D, Ben-Bassat

A, Payne M (2007). Protein engineering of Acidovorax facilis 72W nitrilase for bioprocess

development. Biotechnology and Bioengineering 97: 689-693.

30. Yang F, Zhou Y, Yin L, Zhu G, Liang G, Pu Y (2014). Microcystin-degrading activity of

an indigenous bacterial strain Stenotrophomonas Acidaminiphila MC-LTH2 isolated from

Lake Taihu. PLoS One 9:e86216.

31. Zhang J, Shi H, Liu A, Cao Z, Hao J, Gong R (2015.) Identification of a new microcystin-

degrading bacterium isolated from Lake Chaohu, China. Bulletin of Environmental

Contamination and Toxicology 5:661-666.

Chapter III: Taxonomic and Functional Heterogeneity of Microcystin Degrading Bacteria isolated from Lake Erie Preface

The third chapter for my dissertation has been made into a manuscript for submission to

(Harmful Algae). My co-author Meaghan Balaban has helped in measuring biofilm formation of

MC-degrading isolates and Yuquin Zhang has participated in performing bacterial isolation; Dr.

Youngwoo Seo (University of Toledo) has helped in sampling. My dissertation advisor, Xiaozhen

Mou has advised me extensively with the experimental set up and writing up manuscript.

Abstract

Microcystins (MCs) are among the predominant cyanotoxins in various freshwater environments, including Lake Erie, a Laurentian Great Lake. MCs are chemically refractory and are thought to be primarily degraded by heterotrophic bacteria in natural environments. However, despite the prevalence of MCs in Lake Erie basins, our knowledge about the taxonomic diversity of MC- degrading bacteria in Lake Erie is very limited. The current study obtained thirty-four MC- degrading bacterial pure isolates from Lake Erie surface water. These isolates were characterized taxonomically and examined for their MC-degradation rates under different pH, temperature and with and without the presence of other organic carbon. A dual culture experiment was also performed to examine potential interspecific interactions between MC degraders. Our collection of MC-degrading isolates include similar abundance of gram positive (i.e., Firmicutes

(Brevibacillus and Bacillus) and Actinobacteria (Streptomyces and Microbacteria); 18 isolates)

and gram negative bacteria (i.e., Gammaproteobacteria (Pseudomonas and Stenotrophomonas); 16 isolates). Six of the 34 isolates were motile and had the capacity to form biofilms. The MC- degradation rate of all obtained isolates was impacted by temperature and pH and the presence of cyanobacterial exudates. At optimal pH (7-8) and T (30-35ºC), MC degradation rate of obtained isolates ranged between 0.1-0.25 µg/ml/hr (average 0.2 µg/ml/hr)。 LEw-2010, LEw-24 and LEw-

1002 had the highest MC-degradation rates (0.25 µg/ml/hr ), while LEw-2125, LEw-2082 and

LEw-2079 were found to have the lowest degradation rates (0.1 µg/ml/hr ). Dual culture mixtures showed some isolate combinations to degrade microcystin better at a degradation rate of 0.40

µg/ml/hr as compared to the mono culture average degradation rates of 0.20 µg/ml/hr. These MC- degrading bacteria also show absence of the mlrA gene indicating the absence of the mlr gene based microcystin degradation pathway. Our results suggest a highly diverse microbes that are capable of degrading microcystins are naturally exist in Lake Erie. These highly diverse microbes show great potential to be used in biological treatment of MC-LR contaminated water based on their capacity to form biofilms and degrade microcystin at higher temperatures of 35ºC and alkaline pH of 9. Obtained MC-degrading bacteria had a higher degradation rate compared with isolates previously screened in other studies. The ability for our MC-degrading bacteria to degrade microcystin in the presence of algal exudates and when co cultured with other MC-degrading bacteria, further make them ideal for utilization in biological filters. Absence of the mlrA gene in these MC-degrading bacteria indicates the presence of a novel and uncharacterized microcystin degradation pathway. The results recorded in this study enable us to better comprehend the 60

environmental impact on MC degradation and assess the obtained MC-degraders for their future use in MC bioremediation.

Introduction

Cyanobacterial harmful blooms (CyanoHABs) are common to warm and eutrophic waters and can cause a number of detrimental consequences (Paerl et al., 2013). Harmful consequences of CyanoHABs typically initiate with loss of water clarity, which suppresses the growth of aquatic macrophytes, an important component for fish habitat (Paerl et al., 2013). In addition, large quantity of organic substrates and inorganic nutrients are released during the senescence of blooms, which stimulate growth of aerobic heterotrophic bacteria. This process consumes and even depletes dissolved oxygen and creates hypoxic and anoxic zones in CyanoHAB impacted areas

(Chislock et al., 2013). Moreover, exudates of CyanoHABs often include toxic secondary metabolites with microcystins as the most prevalent type (Codd et a., 1997).

Microcystins (MCs) are a class of monocyclic heptapeptides that are toxic to liver cells

(Pudik et al., 2012). MCs possess a common chemical structure of cyclo-D-Ala1 -X2 -D-MeAsp3

-Z4 -Adda5 -D-Glu6 -Mdha7 (Diehnelt et al., 2006). Over 90 MC isoforms have been identified, among which microcystin-leucine arginine (MC-LR) has leucine (L) at position 2 and arginine (R) at position (4). MC-LR is the most abundant and toxic one among the other isoforms (Dietrich et al., 2005). A number of aquatic organisms like shrimps, molluscs and fish bioaccumulate the toxin

(Paerl et al., 2013).

These toxins are pose as potential carcinogens when they exceed the amounts which have been standardized (Zhang et al., 2017). Therefore, MC-LR concentration in drinking water should not exceed 1 µg/L as recommended by World Health Organization (WHO) (WHO, 1998). Due to 62

the potential hazards posed by the toxin it is essential that we further understand the methods for complete removal of the toxin.

MC-LR and all other MC isoforms possess a monocyclic heptapeptide structure which are non-volatile and hydrophilic in nature, which makes MCs resistant to both physical and chemical breakdowns in natural environment (Sangolkar et al., 2006). MC degradation in natural environments, like lakes and reservoirs which are previously exposed to Cyano HABs, is mainly carried out by the naturally occurring bacteria present in them (Bourne et al., 1994; Krishnan et al., 2018; Mou et al., 2013).

Lakes with a previous history of CyanoHABs are prone to harbor a diversity of bacteria that can degrade MCs and use them as their carbon and/or energy source (Anderson et al., 2002).

Members of Sphingomonaceae family of alphaproteobacteria (such as Sphingomonas, Sphinopyxis and Sphingosinicella) (Bourne et al., 1996; Park et al., 2001; Manage et al., 2009) dominated the pool of MC-degrading bacterial isolates. These bacteria degrade MC-LR using an mlr genes-coded cleavage pathway (Bourne et al., 1996).

Many more other families of MC degraders have been since isolated, such as a number of

Pseudomonas and Stenotrophomonas (gamma-proteobacteria), Acidovorax (beta-protoebcateria) and Bacillus (Firmicutes) (Chen et al., 2010; Krishnan et al., 2018)Many of these new MC- degrading isolates carried no mlr genes (Yang et al., 2014). A metagenomic analysis in Lake Erie suggest a novel non-mlr MC degradation pathway that involved xenobiotic metabolism, however, no direct evidence is available yet (Mou et al., 2013b). 63

A number of factors, like pH, temperature and presence of organic nutrient (nitrate, phosphorous and glucose) are known to affect the rate of microbially mediated MC degradation performed under laboratory-controlled conditions (Schmidt et al., 2014; Li et al., 2011; Ho et al.,

2012). There has also been a study which has utilized immobilized bacteria for the removal of microcystins in a water treatment plant and studied the factors affecting the degradation rate (Tsuji et al., 2006), along with other studies which have summarized the use of naturally occurring MC degrading bacteria in lakes and the factors like pH and temperature affecting them (Gagala and

Mankiewicz-Boczek, 2012).

Lake Erie is the smallest and warmest lake among the five Laurentian Great Lakes

(Hartman et al., 2011) and it has been suffering with a history of eutrophication and sporadic

CyanoHABs (Michalak et al., 2013). In August 2014, a bloom event in the western basin of Lake

Erie led to shut down of water supply for 500,000 residents (Fitzsimmons, 2014). Eradicating MCs from Lake Erie has become a hot research topic. This current chapter describes a continuous and more extensive culturing effort after our initial attempts (Chapter 2) to isolate more microcystin degrading bacterioplankton to enable a clearer view of the diversity of culturable MC-degrading bacteria in Lake Erie. In addition to characterizing taxonomy and phenotypic characteristics, newly obtained MC-degrading bacteria were also examined for their biofilm-forming ability and degradation activities with the presence of algal exudates and other MC-degrading bacteria. The obtained results allowed us to further understand environmental control on MC degradation and evaluate potentials of obtained MC-degraders in MC bioremediation. 64

Methods

Bacterial isolation and screening for MC degrading activities

Samplings were carried out every month from July to October in the years of 2015 and

2016 in the Sandusky Bay and Western Basin of Lake Erie, two areas with frequent and long- lasting CyanoHABs (HAB Bulletin, NOAA, 2017). The method for bacterial isolation and screening was similar to the one in (Chapter 2). Briefly, multiple kinds of solid media were used for isolation of bacteria from Lake Erie and the bacterial cultures were repeatedly streaked to obtain pure culture isolates. These cultures were screened using MT2 Biolog plate assay where the isolates were exposed to MC-LR as the sole carbon source in BG11 medium. Isolates showing an increase in optical density were recorded and confirmed as MC-degrading bacteria.

Phenotypic characterization of individual isolates

The method for phenotypic characterization was similar to the one in (Chapter 2). Briefly, phenotypic characteristics of putative MC-degradation bacterial isolates, including colony pigmentation, cell morphology and gram nature, were examined using standard gram staining procedure (Popescu and Doyle, 2009) and observation of the colony growth on R2A agar

(Ohgiwari et al., 1992). Motility of the strains was identified using the hanging drop technique for motility observation (Clark et al., 1976).

Biofilm formation potential of bacterial isolates was studied by using the assorted media protocol (Fujishige et al., 2006). Briefly, aliquots of 100 µl were taken from fresh cultures of MC- 65

degrading bacteria and transferred into 96-well microtiter plates. After 24 hrs of incubation at 30ºC and in the dark, the microtitre plate was stained with crystal violet for 20 minutes and washed three times with sterile water, followed by a single addition of 95% of ethanol to solubilize the dye.

After discarding the ethanol, absorbance readings were then measured at 570 nm and the biofilm formation for each bacterial isolate was recorded using a Synergy Multimode reader (Biotech, VT,

USA).

Analyses of 16S rRNA and mlrA genes

Paucibacter toxinivorans 16998 and Sphingosinicella microcystinivorans 19791were purchased from DSMZ German Collection of Microorganisms and Cell culture (Braunschweig,

Germany). These two isolates served as positive controls for mlr gene-based MC degradation.

Genomic DNAs of bacterial isolates and positive controls were extracted from each individual isolates using the Ultra Clean Gel Spin DNA Extraction Kit (Mo Bio, CarlsBad, Ca,

USA). Extracted DNAs for isolates was used for the 16S rRNA specific amplification using 27F

(Forward primer) and 1492R (Reverse primer). The PCR amplified region was then examined using 1% agarose gel electrophoresis and the 16S rRNA bands purified using Ultra Clean Gel

Purification kit (Mo Bio, CarlsBad, Ca, USA). The near full length 16S sequence was obtained at

Microgen Corporation, USA. These sequences were then submitted at NCBI under the accession numbers KX185385-KX185419.

Extracted DNA from the MC-degrading bacterial isolates and the positive controls were used for a mlrA based PCR amplification using the primers MF and MR (MF- 5′- 66

GACCCGATGTTCAAGATACT-3′; MR: 5′-CTCCTCCCACAAATCAGGAC-3′) (Yang et al.,

2014) under the following conditions of 95ºC for 3 mins, 95ºC for 1 min, 60ºC for 1 min, 72ºC for 1 min repeated 30 times and 72ºC for 10 mins. The PCR amplified region was then examined using the 1% agarose gel electrophoresis, to confirm the presence of the mlrA region.

Effect of physicochemical variables on MC degradation

Effect of pH and temperature on MC degradation was examined following the same procedures as described in Chapter 2.

Algal exudates were extracted from Microcystis aeruginosa. Briefly, non-toxigenic

Microcystis aeruginosa (ATCC: UTEX LB 2385) were grown in 500ml flasks using BG11 medium under constant illumination at temperature of 27ºC and at 200 rpm (VWR, 1585 shaking incubator). M. aeruginosa cells were harvested when the cells were at their logarithmic phase of growth (about two weeks) by centrifuging 25 ml growth media at 1000 rpm for 10 mins Cell pellets were re-suspended in 10 ml of fresh BG11 medium and sonicated using a Cole Parmer sonicator (Cole Parmer, IL, SA) for 30 sec with pulses every 10 seconds. Sonicated cell suspensions were then centrifuged at 5000 rpm for 15 mins and the supernatant was filtered through 0.2 µm membrane filters. The obtained filtrates were designated as algal exudates and were measured for concentrations of total dissolved organic carbon using a Shimadzu total organic carbon analyzer (Shimadzu, Beachwood, OH, USA).

MC-degrading bacterial isolates (0.1 OD x 2 ml) were grown in test tubes containing MC-

LR (5 mg/ L) and diluted algal exudates (final conc. DOC=12.2 mg/L) in BG11 medium, where 67

the concentration of the DOC used is higher than the concentration typically observed in Lake Erie

(5-10 mg/L). Controls were also established in tubes containing the bacterial suspension in BG11

(3ml) and microcystin (1 mg/L), but not containing the algal exudates. All tubes were incubated at 30ºC and 200 rpm (dark) and 2 ml aliquots were collected from the tubes at intervals of 0, 12 and 24 hrs to track the removal of MC-LR.

Dual culture experiments

Bacteria with MC-LR degradation rates of 0.2 µg/ml/hr or faster were randomly paired for the mixed culture study (Table 5). Bacteria monocultures were harvested at mid-exponential phase and washed with 1× phosphate buffer saline (PBS) three times. The washed cell pellets were re- suspended in BG11 media and the optical density of the culture suspension was adjusted to

OD600- 0.1. Monocultures were organic carbon (OC) starved in new BG11 media for 24 hrs at

30ºC. After the OC starvation, two monocultures were combined (1: 1; v/v) and amended with

MC-LR (1 µg/ml, final concentration) before being incubated at 30ºC and 200-rpm rotation for 48 hrs. No-cell controls were set up and processed with the same procedure. Subsamples were collected at 0, 12, 24 and 48 hrs after start of the incubation.

Microcystin concentration measurement

Microcystins Nodularins ADDA ELISA kit (ABRAXIS Warminster, PA, USA) was used for measuring the MC-LR consumption in samples by using the manufacturer’s protocol. 68

3. Results

3.1. Isolation and characterization of microcystin degrading bacteria

A total of 34 strains were identified as MC-LR degraders based on BIOLOG MT2 assay from 5000 isolates. Following 48 hrs of incubation in MT2 plates, 34 wells with replicates, (i.e.,

34 isolates) containing MC-LR at a concentration of 1 and 10 µg/ml showed significant increases in absorbance values as compared to the no cell controls (P <0.05, Fig 8), reflecting consumption of MC-LR by bacterial isolates. At 0.1 µg/ml of MC-LR, only 7 isolates were positive for consumption of MC-LR within 48 hrs of incubation. These are LEw-24, LEw-1029, LEw-1233,

LEw-1270, LEw-2079, LEw-2123 and LEw-2175.

Obtained MC-degrading bacterial isolates were heterogeneous in their phenotypic traits.

Growing on R2A plates, bacterial colonies appeared white (11 isolates), yellow (13 isolates), orange (6 isolates), pink (3 isolates), and grey (1 isolate). Cellular morphology of these isolates varied among cocci (8 isolates), coccobacilli (11 isolates) and bacilli (15 isolates) (Fig 9). More isolates were gram positive (18) than gram negative (16) (Table 4). Hanging drop method revealed motility of 6 MC-degrading isolates from the 34 isolates. Of the 34 isolates 12 were found to form biofilms (Table 4), among them LEw-24, LEw-2029 and LEw-2082 were also found motile.

Sequences of near full-length 16S rRNA genes revealed that obtained MC-degrading bacteria belonged to the genera of Pseudomonas (6), Stenotrophomonas (10) (Gammaproteobacteria); 69

Bacillus (9), Brevibacillus (1) (Firmicutes); Microbacterium (6), Streptomyces (2)

(Actinobacteria) (Fig 10).

None of the obtained MC-degrading isolates were affiliated with Sphingomonaceae of

Alphaproteobacteria. Moreover, none of these isolates yielded positive results for mlrA gene based

PCR amplification (Fig 11). However, positive controls, i.e., Paucibacter toxinivorans (16998) and Sphingosincella microcystinivorans (19791) did yield positive amplification of mlrA genes.

3.3 Effect of temperature and pH on MC-LR degradation

MC degradation activity was positively correlated with temperatures in the tested range of

15ºC-35ºC for all isolates. Isolates Bacillus cereus LEw-1006, Stenotrophomonas sp. LEw-1029 and Pseudomonas putida LEw-1040 showed positive MC degradation at as low as 15ºC (Fig 12).

At 25ºC, 30ºC and 35ºC, the degradation rate increased and a 100% consumption of MC-LR was observed within 48 hrs of incubation for all isolates (Fig 12). At 25ºC and pH 7, bacterial isolates degraded MC-LR (5 µg/ml) at 0.15-0.20 µg/ml/ while, incubation temperature of 30ºC and 35º showed a degradation rate between 0.20-0.25 µg/ml/hr. The average rate of microcystin degradation was found to be 0.20 µg/ml/hr with highest degradation rates shown by isolates at all tested temperatures of 25 (0.1 µg/ml/hr ), 30 (0.15 µg/ml/hr ) and 35ºC (0.25 µg/ml/hr ).

At 30ºC, bacteria degraded MC-LR the fastest at pH 7 and 8 (Fig 13) with a degradation rate between 0.15-0.20 µg/ml/hr. Lower or higher pH conditions appeared to inhibit MC-LR degradation (P<0.05) for most isolates Exceptions were found for 9 isolates which showed lower

degradation activity at lower pH of 5 and 6 and also at higher pH of 9 (Fig 13). These isolates belonged to Pseudomonas, Bacillus and Stenotrophomonas.

3.4 Effects of algal exudates on MC-LR degradation

Addition of algal exudates to the microcystin degrading isolates showed an increase in the microcystin degradation rate for only one isolate out of the 34 MC-degrading isolates screened.

The isolate Pseudomonas sp. LEw-2029 showed an increase in the MC-LR degradation rate from

0.12 to 0.22 µg/ml/hr after addition of algal exudates; as opposed to Brevibacillus brevis LEw-

2113 which shows a reduction in the degradation rate (Fig 14).The MC-LR degradation rate for the rest of the isolates (32) control and test seem to remain similar with and without addition of the external source of organic C (12.2 mg/L) (Fig 14).

3.5 Dual culture degradation experiment

Degradation rate measured for the mixed culture suspension showed a higher degradation activity for six bacterial combinations when compared with the degradation rate for single culture suspensions (Fig 15). Acidovorax sp. LEw-2 and Stenotrophomonas sp. LEw-2055; along with

B. thuringenesis LEw-2010 and B. pumilus LEw-1282 combinations were observed as having the least degradation rate of 0 when compared to the degradation rate for the single culture suspensions which was between 0.2-0.25 µg/ml/hr. The highest degradation rate was seen for combination of

Pseudomonas sp. LEw-2029 and Bacillus pumilus LEw-1132 (0.4 µg/ml/hr) which had a single culture degradation rate of 0.25 µg/ml/hr. The other isolate combinations which were found to have high degradation rates were Pseudomonas aeruginosa LEw-2010 and Pseudomonas 71

fluorescence LEw-24 (0.35 µg/ml/hr), Microbacterium sp. LEw-2155 and Pseudomonas sp. LEw-

2066 (0.34 µg/ml/hr), Bacillus sp. LEw-1199 and Bacillus altitudinis LEw-1011 (0.3 µg/ml/hr),

B. thuringenesis LEw-2082 and B. pumilus LEw-1270 (0.28 µg/ml/hr). These isolate combinations showed complete consumption of MC-LR within 48 hrs of incubation with single culture degradation rates between 0.2-0.25 µg/ml/hr (Fig 15).

4. Discussion

Previous studies have found Sphingomonadales and mlrA degradation pathway as the primary degradation pathway for complete removal of microcystin (Bourne et al., 1996; Jones et al., 1994). Current work suggests the presence of diverse taxa involved in microcystin degradation and they might adopt a novel degradation pathway (Mou et al., 2013; Chapter 2) as opposed to the mlr gene based degradation pathway (Bao et al., 2016; Kansole et al., 2016; Lezcano et al., 2016).

The 34 MC-degrading isolates were affiliated with, Gamma proteobacteria, Firmicutes and

Actinobacteria (Fig 10). The genera under gamma proteobacteria were Pseudomonas,

Stenotrophomonas. The genera under Firmicutes were Bacillus and Brevibacillus and under

Actinobacteria were Streptomyces and Microbacterium. It was observed that of the 34 isolates 16 belonged to Gamma proteobacteria, 10 belonged to Firmicutes and 8 belonged to Actinobacteria.

Studies on biofilm formation of cultured MC-degrading bacteria (Manage et al., 2009; Yang et al., 2014; Jones et al., 1994) is very limited, although biofilm establishment represents an ideal property for bacteria that can be used to develop “biofilters” for treating MCs in Water Treatment

Plants (WTP). In addition to our studies, only a few other studies have provided indirect evidence 72

of biofilm formation ability for MC degrading bacteria. For example, MC-degrading bacteria have been isolated from sand filters of water treatment plants (Ho et al, 2006; Ho et al., 2007). Due to lack of work in terms of studying the effect of biofilm formation on MC-degrading isolates, our work has highlighted the biofilm formation capacity for some of the isolates. In our study, 3 isolates, i.e., LEw-24, LEw-2055 and LEw-1270, showed both high MC degradation rate (0.25

µg/ml/hr) and strong biofilm formation ability and can potentially serve as good candidate for biofilter development in WTP’s for treatment of MC-contaminated water.

Change in the pH and temperature of the freshwater lake environment results in the formation of harmful cyanobacterial blooms (Paerl et al., 2012). All previously done studies for microcystin degrading bacterial isolates haven’t looked at the effect pH and temperature on microcystin degradation (Zhang et al., 2017; Yang et al., 2014). Most of the isolates showed a higher degradation rate for pH 7 and 8 (Fig 13), but a few isolates did show the presence of degradation activity at pH 9 and 5 with a lower degradation rate. Isolates, such as

Stenotrophomonas sp. LEw-1198 and Stenotrophomonas maltophila LEw-2123, showed degradation activity at alkaline pH (Fig 13) and 35ºC (Fig 12), and they could be potentially used for microcystin degradation in microcystin contaminated lake water due to the alkaline pH and high temperatures in lake waters during the bloom season (Gao et al., 2012).

Degradation with higher efficiency has been shown for various kinds of aromatic compounds when mutually compatible strains are utilized (Babu et al., 1995). In our study, we have noticed that some bacterial combinations are more effective at degradation of microcystin as compared to 73

the others. Degradation activity performed with co-culturing of multiple kinds of bacteria could be made more efficient, if organisms are exposed to bacteria which are commonly encountered in the environment (McDonnell et al., 2015). Microorganisms involved in biological degradation require acclimation to the natural environment and the other microbial populations (Satsangee et al., 1990). Acclimatization is a crucial step in mixed culture degradation of xenobiotic compounds.

It was observed that the combination of P. fluorescence LEw-24 and P. aeruginosa LEw-2010 mixture and Pseudomonas sp. LEw-2029 and Bacillus sp.LEw-1132 were found to have increased degradation rate by almost two folds (Fig 15). Of the combinations that were studied for

MC degradation, LEw-24 + LEw-2010 was found to have the highest degradation rate (0.4

µg/ml/hr). Both the aforementioned isolates, were also shown to be motile and possess the ability to form biofilms which make them potentially useful for development of biofilters in the water treatment plants, as compared to the single cultures which showed a slower degradation rate compared to the mixed culture studies. Concurrent with our observation, Pseudomonas species has also been previously found to possess degradation genes capable of degrading quite a few types of recalcitrant compounds like hydrocarbons, petroleum, oils etc., hence proving its degradation ability (Zhang et al., 1992; Kilbane et al., 1982) and microcystin (Takenaka, 1997).

Bloom affected lakes are abundant in labile organic carbon (Paerl et al., 2011), which may potentially impede the MC degradation (Schimdt et al., 2014). Our study, indicates that addition of an external source of C does not impede the degradation of MC-LR performed by these MC- degrading isolates (Fig 14). With the biofilm formation capacity, mixed culture degradation, 74

degradation in the presence of external source of organic C and the ability to degrade microcystin at an alkaline pH of 9 and high temperature of 35ºC, these bacteria pose as potential candidate for biological degradation of MC-LR in WTP. Further studies are required to evaluate the ability of isolated bacteria to degrade microcystin in a larger scale for water bodies in the environment contaminated with microcystin.

Table 4: Morphological characteristics of microcystin degrading bacteria. Under gram nature, + is gram positive and – indicates gram negative. + and – indicates motile and non motile respectively under motility. ++, + and – under biofilm formation indicates high, moderate and no biofilm formation.

Isolate Species name Color Morphology Gram Motility Biofilm nature (+/-) (++/+/-) (+/-) LEw- 14 Solibacillus silvestris White Cocci + - -

LEw-24 Pseudomonas White dots Cocci - + ++ flourscences

LEw-1002 Bacillus subtilis White Bacilli + - - LEw-1006 Bacillus cereus Pink Bacilli + - -

LEw-1011 Bacillus altitudinis Orange Bacilli + - -

LEw-1029 Stenotrophomas sp CV67 White Bacilli - - +

LEw-1040 Pseudomonas putida Dark pink Cocci - - -

LEw-1049 Stenotrophomonas White Coocobacilli - - - maltophila

LEw-1052 Pseudomonas FB44 Beige Cocci - - -

LEw-1055 Microbacterium ZT6 Brown Bacilli + - -

LEw-1069 Stenotrophomomas CV67 Grey Cocci - - ++

LEw-1072 Bacillus cereus Orange Bacilli + - +

LEw-1132 Bacillus pumilus White Bacilli + + -

LEw-1198 Stenotrophomomans sp Pink Coccobacilli - - - PA44 76

LEw-1199 Bacillus aerophilus Opaque Bacilli + - ++

LEw-1209 Micromonospora sp C7 Yellow Bacilli + - -

LEw-1232 Microbacterium spTHG White Bacilli + + -

LEw-1233 Bacillus pumilus White Cocci + + -

LEw-1270 Bacillus pumilus White Bacilli + - ++

LEw-1278 Stenotrophomonas Yellow Cocci - - - maltophila

LEw-1282 Bacillus pumilus White Bacilli + + -

LEw-1304 Microbacterium spB015 Light Bacilli + + + yellow LEw-2009 Srenotrophomonas White Coccobacilli - - - maltophila LEw-2010 Pseudomonas aeruginosa White Cocci - + + LEw-2029 Pseudomonas sp FBF35 Orange Coccobacilli - - ++

LEw-2055 Streptomyces flavoviridis Grey Coccobacilli + - ++

LEw-2079 Stenotrophomonas Light Cocci - - - maltophila yellow

LEw-2082 Bacillus thuringeneisis Yellow Bacilli + + ++

LEw-2087 Bacills subtilis White Cocci + + LEw-2089 Stenotrophomonas Orange Bacilli - - - rhizophila

LEw-2113 Brevibacilllus brevis Beige Coccobacilli + - + LEw-2123 Stenotrophomonas Orange Bacilli - - - maltophila

LEw-2125 White Coccobacilli + - -

LEw-2174 Stenotrophomonas sp Orange Coccobacilli - - + PA44

LEw-2175 Stenotrophomonas Bright pink Bacilli - - - maltophila

Table 5: Combination of the MC-degrading isolates used for mixed culture degradation

Mixed culture isolates Identity

LEw (2+2055) Acidovorax facilis & Pseudomonas sp.

LEw(2155+2166) Microbacterium sp & Pseudomonas sp.

LEw (2029+1132) Pseudomonas sp & Bacillus sp

LEw(1232+2175) Microbacterium sp & S. maltophila

LEw(1199+1011) Bacillus sp. & Bacillus altitudinis

LEw(2082+1270) B. thuringenesis & B. pumilus

LEw(1029+2089) S. rhizophila& S.rhizophila

BIOLOG MT2 3 LEw(2010+24) B. thuringenesis & P. fluorescence

2.5 LEw(2010+1282) B. thuringenesis & B. pumilus

2 1

1.5

0.5 Optical density (630 nm) (630 densityOptical 0

LEw-24

LEw-14

LEw-1055 LEw-1232 LEw-1006 LEw-1011 LEw-1029 LEw-1040 LEw-2087 LEw-1049 LEw-1052 LEw-1069 LEw-1072 LEw-1132 LEw-1198 LEw-1199 LEw-1209 LEw-1233 LEw-1270 LEw-1282 LEw-1304 LEw-2029 LEw-2055 LEw-2079 LEw-2082 LEw-2089 LEw-2113 LEw-2123 LEw-2174 LEw-2175 LEw-1002 Isolates 0 0.1 1 10 µg/ml

Fig 8: Microcystin degradation of the MC-degrading isolates as seen in the BIOLOG MT2 screening. Bacterial isolates which showed a significant increase in their optical density for MC-LR concentrations of 0.1, 1 and 10 g/ml were depicted in this bar graph. µ

a) b) c)

) )

Fig 9: Microscopic observation for gram staining performed for MC-degrading bacterial isolates with morphologies of bacilli (a)cocci (b) and coccobacilli (c) . Magnification used for observing the stains was 1000× Scale bar 0.2 µm

Gamma proteobacteria

Actinobacteria

Firmicutes

Fig 10: A neighbor-joining phylogenetic tree based on partial sequence of 16S rRNA gene showing the relationship sequences of isolated MC-degrading bacteria and their closely related relatives. The GenBank accession numbers of the sequences are shown in parentheses. Bootstrap values that are higher than 50% are show at the branch nodes (1,000 resampling). The scale bar represents 0.02 nucleotide substitutions per position.

+ve ctrl MC +

16998 +ve 19791 ctrl 24 2010 1033

+ve ctrl 750 650

400

200

mlrA mlrA

Fig 11: Gel image for the mlrA based amplification performed for MC-degrading isolates (LEw-24, LEw-2 and LEw-1030). mlrA amplification was observed for the positive (+ve) control Sphingosinicella microcystinivorans (16998) and Paucibacter toxinivorans (19791)

Fig 12: Effect of temperature on MC-LR degradation rate as shown in a heat map. The gradient of colors from blue to red indicate the increasing degradation rate (µg/ml/hr) for MC- degrading isolates.

Fig 13: Effect of pH on MC-LR degradation rate as shown in a heat map. The gradient of colors from blue to red indicate the increasing degradation rate (µg/ml/hr) for MC-degrading isolates.

Fig 14: Heat map representing the degradation rates (µg/ml/hr) for isolates when incubated in a suspension containing other organic nutrients. The gradient of colors from blue to red represent the degradation rates of the isolates. Control represents the degradation rate of the isolates when incubated in a suspension containing only MC-LR and test represents the microcystin degradation rate for the isolates when incubated in suspension containing MC-LR and organic C source.

0.45 0.4 0.35 0.3 0.25 0.2 0.15 0.1

Degradation rate (µg/ml/hr) rate Degradation 0.05

LEw-2

LEw-24

LEw-2055 LEw-2155 LEw-2066 LEw-2029 LEw-1132 LEw-1233 LEw-2175 LEw-1199 LEw-1011 LEw-2082 LEw-1270 LEw-1029 LEw-2089 LEw-2010 LEw-2010 LEw-1282

LEw-(2+2055)

LEw-(2010+24)

LEw-1132+2029)

LEw-(2155+2066) LEw-(1233+2175) LEw-(1011+1199) LEw-(2082+1270) LEw-(1029+2089) LEw-(2010+1282) Isolates

Fig 15: MC-LR degradation rates (µg/ml/hr) as measured for mixed culture* MC- degrading isolates. Degradation rates were compared between the individual MC- degrading isolates and when in mixture has been added for the isolate combinations which have a higher degradation rates than the individual isolates.

References 1. Amman, R., Binder, B., Olson, R., Chisholm, S., Devereux, R and Stahl, D (1990).

Combination of 16S rRNA-Targeted Oligonucleotide Probes with Flow Cytometry for

Analyzing Mixed Microbial Populations. Applied and Environmental Microbiology 56(6):

1919-1925.

2. Anderson, D., Gilbert, P and Burkholder, J (2002). Harmful algal blooms and

eutrophication: Nutrient sources, composition, and consequences. Estuaries 25(4):704-

726.

3. Babu, K., Ajith-Kumar, P and Kunhi, A (1995). Simultaneous degradation of 3-

chlorobenzoate and phenolic compounds by a defined mixed culture of Pseudomonas spp.

World Journal of Microbiology & Biotechnology 11: 148-152.

4. Bao, Z and Wu, Y (2016). Biodegradation of microcystin-LR by an amino acid-degrading

anaerobic bacterium. Desalination and Water Treatment 57 (2):870-880.

5. Bourne, D., Jones, G., Blakeley, R and Jones, A (1996). Enzymatic pathway for the

bacterial degradation of the cyanobacterial cyclic peptide toxin microcystin LR.. American

Society of Microbiology 62 (11):4086-4094.

6. Bourne, D., Riddles, P., Jones, G., Smith, W and Blakeley, R (2001). Characterisation of a

gene cluster involved in bacterial degradation of the cyanobacterial toxin microcystin LR.

Environmental Toxicology 16(6): 523-534.

7. Chen, J.; Hu, L.B.; Zhou, W.; Yan, S.H.; Yang, J.D.; Xue, Y.F.; Shi, Z.Q. Degradation of

Microcystin-LR and RR by a Stenotrophomonas sp. strain EMS isolated from Lake Taihu,

China. Int. J. Mol. Sci. 2010, 11, 896–911

8. Chislock, . F., Doster, E., Zitomer, R and Wilson, A. (2013). Eutrophication: Causes,

Consequences, and Controls in Aquatic Ecosystems. Nature Education Knowledge

4(4):10-20.

9. Christoffersen, K., Lyck, S and Winding, A (2002). Microbial activity and bacterial

community structure during degradation of microcystins. Aquatic Microbial Ecology

27(2):125-136.

10. Clark W (1976) A simplified Leifson flagella stain. J Cli. Microbiol 3: 632-634.

11. Codd G.A., Ward C.J., Bell S.G. (1997) Cyanobacterial Toxins: Occurrence, Modes of

Action, Health Effects and Exposure Routes. In: Seiler J.P., Vilanova E. (eds) Applied

Toxicology: Approaches Through Basic Science. Archives of Toxicology

(Supplement), vol 19. Springer, Berlin, Heidelberg

12. Dietrich, D and Hoeger, S (2005). Guidance values for microcystins in water and

cyanobacterial supplement products (blue-green algal supplements): a reasonable or

misguided approach? Toxicology and Applied Pharmacology 203(3): 273-289.

13. Diehnelt CW, Dugan NR, Peterman SM and Budde WL (2006). Identification of

microcystin toxins from a strain of Microcystis aeruginosa by liquid chromatography

introduction into a hybrid linear ion trap-Fourier transform ion cyclotron resonance mass

spectrometer. Analytical Chemistry 78:501–512.

14. Fitzsimmons, E (2014). Tap Water Ban for Toledo Residents. New York Times A:12.

15. Fujishige, N., Kapadia, N., DeHoff, P and Hirsch, A (2006). Investigations

of Rhizobium biofilm formation. FEMS Microbiology Ecology 56 (2):195-206.

16. Gagala, I and Mankiewicz-Boczek, J (2012). The natural degradation of microcystins

(cyanobacterial hepatotoxins) in freshwater-the future of modern treatment systems and

water quality improvements. Polish Journal of Environmental Studies 21(5):1125-

1139.

17. Gao, Y., Cornwell, J., Stoecker, D and Owens, M (2012). Effects of cyanobacterial-

driven pH increases on sediment nutrient fluxes and coupled nitrification-

denitrification in a shallow fresh water estuary. Biogeosciences 9: 2697-2710.

18. Giaramida, L., Manage, P., Edwards, C., Singh, B and Lawtin, L., (2013). Bacterial

communities response to microcystins exposure and nutrient availability: Linking

degradation capacity to community structure. International Biodeterioration &

Biodegradation 84(2013):111-117.

19. Hartmann, H (1990). Climate change impacts on Laurentian Great Lakes levels.

Climatic Change 17(1):49-67.

20. Ho, L., Meyn, T., Keegan, A., Hoefel, D and Newcombe, G (2006). Bacterial

degradation of microcystin toxins within a biologically active sand filter. Water

Research 40(4):768-774.

21. Ho, L., Hoefel, D., Saint, C and Newcombe, G (2007). Isolation and identification of a

novel microcystin-degrading bacterium from a biological sand filter. Water Research

41(20): 4685-4695.

22. Ho, L., Tang, T., Monis, P and Hoefel, D., (2012). Biodegradation of multiple

cyanobacterial metabolites in drinking water supplies. Chemosphere 87(10):1149-

1154.

23. Jones G, Orr P (1994) Release and degradation of microcystin following algicide treatment

of a Microcystis aeruginosa bloom in a recreational lake, as determined by HPLC and

protein phosphatase inhibition assay. Water Res 28:871-876

24. Kilbane, J., Chatterjee, D., Karns, J., Kellogg, S and Chakrabarty, A (1982).

Biodegradation of 2, 4, 5-Trichlorophenoxyacetic Acid by a Pure Culture of

Pseudomonas cepacia. Applied and Environmental Microbiology 44(1): 72-78.

25. Krishnan, A., Zhang, Y and Mou, X., 2018. Isolation and characterization of microcystin-

degrading bacteria from Lake Erie. Bulletin of Environmental Contamination and

Toxicology. 101(5):617-623.

26. Lezcano, M., Moron-Lopez, J., Agha, R., Lopez-Heras, I., Nozal, L., Quesada, A and

El-Shehhawy, R (2016). Presence or absence of mlr genes and nutrient concentrations 90

co-determine the microcystin biodegradation efficiency of a natural bacterial

community. Toxins 8(11): 1-17.

27. Li, J., Shimizu, K., Sakharkar, M., Utsumi, M., Zahng,Z., and Sugira, N (2011).

Comparative study for the effects of variable nutrient conditions on the

biodegradation of microcystin-LR and concurrent dynamics in microcystin-degrading

gene abundance. Bioresource Technology 102(20):9509-9517.

28. Manage P, Edwards C, Singh B, Lawton L (2009) Isolation and identification of novel

microcystin-degrading bacteria. Appl Environ Microbiol 75:6924–6928.

29. McDonnell, M and Hahs, A (2015). Adaptation and Adaptedness of Organisms to

Urban Environments. Annual Reviews of Ecology, Evolution, and Systematics 46: 261-

280.

30. Michalak, A., Anderson, E., Beletsky, D., Boland, S., Bosch, N., Brdigemena, T.,

Chaffin, J., Cho, K., Confesor, K., Daloglu, I., DePinto, J., Evans, M., Fahnenstiel, G.,

He, L., Ho, J., Johengen, T., Kuo, K., LaPorte, E., Liu, X., McWilliams, M., Moore,

M., Posselt, D., Richards, R., Scavia, D., Steiner, A., Verhamme, E and Zagorski, M

(2013). Proceedings of the National Academy of Sciences of the United States of

America 1(1):1-15.

31. Mou X, Jacob J, Lu X, Robbins S, Sun S, Ortiz, J (2013a). Diversity and distribution of

free-living and particle-associated bacterioplankton in Sandusky Bay and adjacent waters

of Lake Erie Western Basin. Journal of Great Lakes Research, 39: 352-357. 91

32. Mou X, Lu X, Jacob J, Heath R (2013b) Metagenomic identification of bacterioplankton

taxa and pathways involved in microcystin degradation in Lake Erie. PLOS ONE 8:1-

14.e61890.

33. Ohigawari, M., Matsushita, M and Matsuyama, T (1992). Morphological changes in

growth phenomena of bacterial colony patterns. Journal of Physical Scoiety of Japa

61(7):816—822.

34. Paerl, H., Xu, H., McCarthy, M., Zhu, G., Qin, B., Li, Y., Gardner, W (2011).

Controlling harmful cyanobacterial blooms in a hyper-eutrophic lake (Lake Taihu,

China): The need for a dual nutrient (N & P) management strategy. Water Research

45(5):1973-1983.

35. Park, H., Sasaki, Y., Maruyama, T., Yanagisawa, E., Hirashi, A., Kato, K (2001).

Degradation of the cyanobacterial hepatotoxin microcystin by a new bacterium isolated

from a hypertrophic lake. Environmental Toxicology 1(1):337-344.

36. Paerl, H and Paul, V (2012). Climate change: Links to global expansion of harmful

cyanobacteria. Water Research 46(5): 1349-1363.

37. Paerl, H., Fulton, R., Moisander, P and Dyble, J (2001). Harmful Freshwater Algal

Blooms, With an Emphasis on Cyanobacteria. The Scientific World Journal 1: 76-113.

38. Popescu, A and Doyle, R (1996). The gram stain after more than a century. Biotechnic

and Histochemistry 71 (3): 145-151.

39. Pudik, J., Prinsep, M., Wood, S., Kaufononga, S., Cary, S and Hamilton, D (2014).

High Levels of Structural Diversity Observed in Microcystins from Microcystis

CAWBG11 and Characterization of Six New Microcystin Congeners. Marine Druga

12(11):5372-5395.

40. Sangolkar, L., Maske, S and Chakrabarti, T (2006). Methods for determining

microcystins (peptide hepatotoxins) and microcystin-producing cyanobacteria. Water

Research 40(19):3485-3496.

41. Satsangee, R and Ghosh, P (1990). Anaerobic degradation of phenol using an

acclimated mixed culture. Applied Microbiology and Biotechnology. 34(1): 127-130.

42. Schmidt, J., Wilhelm, S and Boyer, G (2014). The Fate of Microcystins in the

Environment and Challenges for Monitoring. Toxins 6:3354-3387.

43. Takenaka, S and Watanabe, M (1997). Microcystin LR degradation by Pseudomonas

aeruginosa alkaline protease. Chemosphere 34(4):749-757.

44. Tsuji, K., Asakawa, M., Anzai, Y., Sumino, T and Harada, K (2006). Degradation of

microcystins using immobilized microorganism isolated in an eutrophic lake.

Chemosphere 65(2006):117-124.

45. WHO (1998) Guidlines for Drinking-Water Quality, 2nd edition. Geneva: World Health

Organization. 95–110

46. Yang, F., Zhou, Y., Yin, L., Zhu, G., Liang, G and Pu, Y (2014). Microcystin-Degrading

Activity of an Indigenous Bacterial Strain Stenotrophomonas acidaminiphila MC-LTH2

Isolated from Lake Taihu. PLOS One 9(1): 1-7.

47. Zhang, J., Lu, Q., Ding, Q., Yin, L and Pu, Y (2017). A Novel and Native Microcystin-

Degrading Bacterium of Sphingopyxis sp. Isolated from Lake Taihu. International

Journal of Environmental Research and Public Health 14(10): 1-14.

Chapter IV: Identification of novel microcystin degrading pathway adopted by bacterioplankton from Lake Erie using transposon mutagenesis and LC-MS

Abstract

Bacterial community coexisting with the toxic cyanobacterial blooms in Lake Erie have been shown to possess the ability to degrade microcystin (MC) (Chapter 2 and 3). Among the MC- degrading bacteria identified, Pseudomonas fluorescence LEw-24 displayed the capacity to degrade microcystin at a higher degradation rate of 0.25 µg/ml/hr and showed the absence of mlrA gene. Therefore, LEw-24 was used for random transposon mutagenesis performed using Tn5 transposon. This mutation was performed to mutate the region of MC-LR degradation in bacteria

Pseudomonas fluorescence LEw-24. Mutants obtained were then screened for the presence of MC-

( MC-non degrading) mutant using MT2 Biolog assay as seen in Chapter 2&3. The mutated region was sequenced and identified using Sanger sequencing. To examine potential products and intermediates of MC degradation, aliquots from LEw-24 cell extract and MC-LR mixture were taken at intervals and HPLC/MS analysis performed to identify the degradation intermediates. The transposon mutagenesis yielded 15000 mutants which were screened and the presence of five MC- mutants were identified. The sequencing results for the mutants revealed the genes to be present adjacent to 16S rRNA genes for Pseudomonas along with some unidentified protein based genes.

Potential intermediate peaks having m/z ratios of 1061 and 509.2 were recorded in the HPLC/MS

data. Future prospects include identifying the other genes and intermediates involved in microcystin degradation for bacteria from Lake Erie.

1 Introduction

Microcystins (MCs) are a class of cyclic heptapeptide toxin which are most prevalent among the cyanotoxins commonly observed in lakes affected with harmful cyanobacterial blooms

(Dietrich et al., 2005). These toxins are tumor promoting hazardous toxins which causes symptoms of nausea, vomiting and diarrhea on consumption (Falconer et al., 1992). All MCs have a common structure of cyclo-D-Ala1-X2-D-MeAsp3-Z4-Adda5-D-Glu6-Mdha7. MC-LR is identified as a variant of MCs which has leucine (L) at positon 2 and arginine (R) at position 4, and is also considered as the most toxic among the variants of MCs commonly observed (Fontanillo et al., 2018).

The degradative pathway adopted by MC-degrading bacteria has been firstly identified in

Sphingomonas sp. ACM-3962 as an mlr gene based step-wise cleavage process (Fig 2; Jones et al., 1994). MC-LR posseses a cyclic heptapeptide structure (Sangolkar et al., 2006) where the enzyme MlrA (microcystinase) initiates the degradation by breaking up the ring and linearizing the cyclic MC-LR. The linearized MC-LR has much lower toxicity than cyclic MC-LR (Jones et al., 1994). Next, a serine peptidase (MlrB) catalyzes the linearized MC-LR at the Ala-Leu peptide bond and produce a tetrapeptide. Finally, the third enzyme (MlrC), cuts the peptide bonds randomly resulting in undetectable peptide fragments and amino acids. Since the initial cleavage

is the most critical step in MC-LR degradation, so far genetic analyses of MC-LR degradation

have been mostly based on analysis of mlrA gens (Jones et al., 1994).

Sphingosinicella, Sphingopyxis, Sphingomonas, and Novosphingobium belonging to the family

Sphingomonadaceae have been shown to possess the mlrA genes (Bourne et al., 1996; Park et al.,

2001; Kormas et al., 2013).

Studies have shown the absence of mlrA gene in microcystin degrading bacteria using PCR

based assays (Manage et al., 2010; Yang et al., 2014). Absence of mlrA gene suggests two

possibilities that, either the mlrA gene are specific for the genus Sphingomonas or, the MC-

degrading bacteria adopts a novel degradation pathway.

Metagenomic study supporting the second hypothesis have recorded low occurrence of mlr genes in bacterioplankton from Lake Erie (Mou et al., 2013). It was also seen that the abundance of mlr gene did not show a change on addition of MC-LR to the bacterioplankton (Mou et al., 2013). A greater presence of xenobiotic metabolism related genes was also recorded for bacterioplankton in Lake Erie (Mou et al., 2013). Xenobiotic metabolism is widely spread among the different domains of life and has shown its role in MC degradation in aquatic eukaryotes

(Peregrin-Alvarez et al., 2009).

In my previous studies (Chapter 2&3) MC-degrading bacteria from Lake Erie were

found to show absence of mlrA genes indicating the presence of a novel and uncharacterized

pathway. In this research we hypothesized that xenobiotic metabolic pathway is present in

microcystin degrading bacteria from Lake Erie the presence of an unknown metabolic pathway present in microcystin degrading bacteria from Lake Erie.

In the current study we have used isolate ID Pseudomonas fluorescence LEw-24 and identified the presence of novel microcystin degrading gene and intermediates. MC-LR degradation rate for the isolate LEw-24 was found to be higher compared with the other MC- degrading bacteria. Since this bacteria belongs to the genus Pseudomonas which possesses the ability to degrade multiple recalcitrant compounds (Barathi et al., 2001) followed by its high MC- degradation rate of 0.2ug/ml/hr, makes this bacteria a potential model for identification of the novel microcystin degrading pathway.

2 Materials and methods

2.1 Bacterial isolates

Pseudomonas flourscences LEw-24 was grown in 5 ml of LB (Luria Bertani: Tryptone:

10g/L, Yeast extract: 10g/L, NaCl: 5g/L, pH 7) media for 24-48 hrs. The culture was diluted in 20 ml of LB media in a 200ml conical flask such that the Optical density (OD600) of the incubated culture is 0.2-0.5. These cells were used for preparing the chemically competent cells.

2.2 Preparing chemically competent cells

The diluted culture (prepared above) (10ml) was poured into two pre-chilled falcon tubes and incubated in ice for 10 minutes and then centrifuged at 300 rpm for 10 mins at 4ºC. Cell pellet was suspended in transformation storage solution (TSS) (Epibio, MI, USA) and 100 µl of cell suspension was taken and transferred to chilled small Eppendorf tubes. The Eppendorf tubes were then immediately placed at -80ºC for 10 mins. These chemical competent cells were then used for transposon mutagenesis.

2.3 Tn5 transposon mutagenesis

An Ez Tn5 (Kanr) transposon mutagenesis kit (Epibio, Madison, WI, USA) was used for performing the random mutagenesis on the chemically competent cells prepared above. A transposome mixture containing the transposon (1 µl) along with the transposase (1µl) and the chemically competent MC-degrading cells (500µl) was prepared. This transposome mixture was used for mutation of the cells according to the instructions on the kit. After incubation, the mutated

cell suspension was plated on LB plates containing kanamycin (100 µg/ml) and incubated at 37ºC for 24 hrs in the dark.

Kanamycin resistant strains which were able to grow on LB Kan+ plates were isolated using a tooth pick for MT2 Biolog plate analysis. The Tn5 mutated cells were grown in LB liquid media

(5ml) for 24 hrs at 30ºC under 200 rpm. The mutants grown in suspension were screened using the

MT2 Biolog assay protocol similar to the one used in Krishnan et al. (2018) for identifying the presence of the MC- (MC non degrading) isolate among theTn5 transposon mutants. In the screening process, mutants were inoculated in liquid LB media and the culture suspension were centrifuged at 2000 rpm for 10 mins and pelleted cells were washed thrice with 1×PBS (5 ml).These washed cells were then re-suspended in BG11 and incubated without carbon for 24 hrs.

These C starved cells were added to the wells in BIOLOG MT2 plates along with MC-LR

(1µg/ml). The MT2 plates were incubated at 30ºC for 24 hrs and the optical density for the wells were recorded at intervals of 12 hrs at 630nm. The isolates showing a decrease in optical density was recorded as a MC- mutants.

2.4 Restriction endonuclease cleavage analysis and growth assay

Putative MC- mutants were grown in liquid LB media (30ºC, 200rpm, 24 hrs) and the cell pellet used for DNA extraction. Total DNAs from obtained MC- mutants were isolated using the

Charge switch gDNA mini bacterial extraction kit (Thermo Fisher Scientific, CA, USA) following the protocol on the kit. Extracted genomic DNAs were stored at 4oC and cleaved at the mutated region using restriction enzyme specific for the Tn5 mutant. 100

Transposon Tn5 contains two Hind III cleavage sites and a single Bam HI site. These enzymes (Invitrogen, USA) were used to locate the sites in Tn5 transconjugants prepared in the procedure above (Tn5 mutagenesis) where the restriction endonuclease digested DNA was utilized for performing gel electrophoresis using 1% agarose gel at 100 volts for 30 mins. The bands obtained were cut, and purified using the Ultra Clean Gel Spin DNA extraction kit (Mo Bio

Carlsbad, California, USA). The extracted DNA was sequenced using Sanger sequencing and the sequences obtained were run in NCBI database using BLASTx to identify for matches of proteins.

Mutant (A45, D1, F79, F82 and F72) and wild type cultures were grown in LB medium (5 ml) and incubated at 30ºC for 24 hrs under 200 rpm. Optical density measurements were taken at 600 nm using an UV/Vis Spectrophotometer (Beckman Coulter Inc, Brea, CA, USA) for the wild type and mutant cultures at intervals of 0, 12, 24 and 48 hrs. These optical density measurements were used to prepare a growth curve for the mutant and wild type isolates in order to confirm the mutation of

16S rRNA gene in the Tn5 mutants.

2.5 Enzyme activity assay

Cultures of Pseudomonas flourscences LEw-24 were inoculated in LB medium (10 ml) and incubated overnight at 30º C and 200 rpm for 24 hrs. The cultures were harvested after 24 hrs and centrifuged at 2000 rpm for 10 mins to obtain a pellet, the supernatant was decanted, and an equal volume (5ml) of PBS added. These re-suspended cells were re-pelleted, by centrifugation at

2000 rpm for 10 mins and the supernatant was discarded, this was repeated three times. The re- 101

suspended cells were sonicated for 30 seconds using a Cole Palmer 8894 sonicator (Cole Palmer,

IL, and USA). The CE (cell free extract) in the supernatant was poured out and filtered and used for the enzyme activity assays. Enzyme assays were performed by using 0.25 ml of CE and microcystin LR at final concentration of 1µg/ml, and PBS was used to make the assay volume upto 1.5 ml. All assay mixtures were prepared in 2-ml Eppendorf tubes on ice, vortexed and incubated at 30ºC. Samples (200 ml) were removed at 0, 6, 8 and 10 hrs after incubation and centrifuged at 15,000 rpm for 5 min.

2.6 High performance liquid chromatography (HPLC)

MC-LR standard and incubated CE were analyzed by a reverse-phase HPLC (Agilent mass

HP 1100 HPLC, Agilent, Santa Clara, CA). This detector was equipped with a UV detector set at

238nm for detection, and a Spherisorb ODS-2 C18 column (Agilent Technologies, Santa Clara,

CA) (5 mm; 250 by 4.6 mm) for separation of the compounds. Volume of injected samples were

100µl (Shamsollahi et al., 2014) and the column temperature was set at 30ºC. All reagents that were used were HPLC grade. Two mobile phases were used, namely: A) 32 % acetonitrile B)

55% acetonitrile. A gradient run was performed using these isolates with the following protocol:

100% A at 0 min, 50% A at 12 min, 100% B at 15 mins, 100% A at 25 mins and 100% B at 60 mins. The flow rate used for the method was 1 ml/min (Shamsollahi et al., 2014).

2.7 Ion spray mass spectrometry (MS)

A triple-quadrupole mass spectrometer (Bruker Esquire 3000plusMA, USA) was used to identify the degradation products and intermediates present in the aliquots obtained from the above 102

process. A pneumatically assisted electrospray (ion spray) interface was used for sample introduction into the atmospheric pressure ionization source. Ion spray MS was performed for identification of degradation products. Reverse HPLC was used to purify and separate the degradation products and then identified based on their mass/charge (m/z) ratio using MS.

3 Results

3.1 Mutant screening and Sanger sequencing

A mutant library of 15,000 mutants was created and screened for the presence of the MC- mutant that lost their ability of MC degradation. The screening process yielded five MC- mutants

(A45, D1, F79, F82, and F72). These mutants had a significantly lower optical density than the other mutants signifying mutation of the microcystin degrading gene (Fig 16). Positive control performed for the isolates showed a significant increase in the optical density indicating the degradation of microcystin.

Sanger sequencing performed for the mutated region yielded 1300 bp region for all the five mutants observed. On performing BLAST the region was found to be similar (97%) to the 16S rRNA region of Pseudomonas fluorescence (Table 6). It was observed that the mutants were shown

103

to grow slowly as compared to the wild type (Fig 17). This suggests Tn5 based random mutation of one of the 16S rRNA regions present in Pseudomonas fluorescens LEw-24.

3.2 Enzyme assay-HPLC

Aliquots collected at various time points were characterized using HPLC. MC-LR was eluted out at 2.1 mins followed by the elution of degradation product A1 and B1 at 1.9 and 2.5 mins respectively (Fig 18). Degradation product A1 appears after 6 hours of incubation, followed by the elution of degradation product B1 after 8 hrs of incubation. Complete removal of MC-LR was observed after 10 hrs of incubation followed by disappearance of degradation product A1 (Fig 18).

Degradation product B1 was still found in the suspension after complete disappearance of MC-

LR.

3.3 Identification of degradation of products

Degradation product A1 was identified with an m/z ratio of 509.2 (Fig 18) and degradation product B1 was identified as the product with an m/z ratio of 1061.

MS-MS analysis was performed for further identification of the MC-LR degradation product. The degradation products didn’t produce a match to the standard breakdown products of

MC-LR via MS (Puddick et al., 2015). With multiple attempts we were unable to identify the exact chemical structure of the compounds identified in the MS analysis. Further work will be required 104

for identification of the compounds using MS/MS. The degradation products which have been identified for the mlr based degradation pathway (Bourne et al., 1996) did not bear a similarity to the degradation products identified in our study. This signifies the presence of a novel microcystin degradation pathway with novel genes and degradation intermediates.

Discussion

Microcystin degradation mediated by bacterial cultures obtained from various lakes around the world has been studied for their degradation properties (Somdee et al., 2012; Manage et al.,

2009; Yang et al., 2014), but little to no work has been done looking at the novel microcystin degrading pathway. mlr cluster genes (mlrABCD), intermediates and products involved in microcystin degradation was first identified for the Sphingomonadales strain (Bourne et al., 1996).

In my previously done work we have found MC-degrading bacteria belonging to Actinobacteria,

Firmicutes, Proteobacteria of the alpha and gamma sub –divisions (Chapter 2&3; Krishnan et al.,

2018). These bacteria showed the absence of mlr cluster genes after a mlrA based amplification. mlrA gene is the first gene responsible for cleaving the cyclic ring structure of MC-LR, reducing the toxicity of MC-LR by 180 times (Jones et al., 1994). Previous studies performed with the metagenomic identification for bacteria from Lake Erie (Mou et al., 2013) has observed the abundance of xenobiotic metabolism genes as opposed to the mlr cluster of genes.

Previously identified degradation pathway showed the presence of degradation product and

B, of which A is the linearized product of MC-LR- NH2–Adda–Glu(iso)–methyldehydroalanine– 105

Ala–Leu–b-methylaspartate–Arg–OH (1013) and the other is a tetrapeptide NH2-Adda-Glu(iso)- methyldehydroalanine-Ala-OH (Bourne et al., 1994). In the current work, we identified the presence of two degradation products A1 and B1. The m/z ratio of degradation product A was identified as 1061 and that of B was 509.2 (Fig 18). These products remain unidentified in terms of the structure, since they are novel products not observed earlier for microcystin degradation.

The possibility of 16S rRNA gene being involved in microcystin degradation has been a novel finding in this study (Table 1). After multiple sequencing processes we identified the presence of microcystin degradation gene to be potentially present in the 16 S rRNA region of the bacteria, but further studies are required to confirm the results. Previous studies have attributed only taxonomic identification to 16S rRNA gene (Janda et al., 2007), but the current work shows the possibility of degradation to be lying in the 16 S rRNA gene which would be one of the unique findings for this chapter.

The presence of novel degradation products and novel genes seem to be in concurrence with the study proposing the presence of an alternate degradation pathway in bacteria from Lake Erie

(Mou et al., 2013).

Further confirmation for the mutation of the degradation gene was obtained by performing a comparative study for microcystin degradation in wild type vs mutant culture. Wild type culture showed an increase in the optical density when incubated in culture suspension containing MC- 106

LR, as opposed to the mutant which showed a decrease in the optical density when incubated in suspension containing MC-LR as the sole carbon source (data not shown). This was in concurrence with the screening process performed earlier for identification of MC- mutant

This work has highlighted the presence of alternate microcystin degradation pathway present in bacteria from Lake Erie by identifying the degradation intermediates and genes involved in the pathway.

Conclusion

Microcystin degrading bacteria in Lake Erie adopts a unique and novel degradation pathway as compared to the other microcystin degrading strains studied until now. Complete degradation of the toxin was observed within 8 hrs of incubation with the toxin. Potential degradation products

A1 and B1 were identified as possessing the m/z ratio 509.2 and 1061 respectively. Degradation product B1 was found to remain in the suspension after complete disappearance of MC-LR.

Multiple confirmatory tests looking at the presence and absence of degradation in mutant vs wild type bacteria further help us speculate the involvement of 16S rRNA gene in the microcystin degradation of the bacteria

107

0.6 MUTANT SCREENING 0.5

0.4

0.3

0.2

0.1 Optical density (630 (630 nm) density Optical

0 Wild A10 B3 C2 F7 A45 D1 F82 F79 F 72 type

Isolates 0 hr 24 hr 48 hr

Fig 16: MC-LR degradation 108of the Tn5 mutants was identified using absorbance measurements in BIOLOG MT2 screening

Mutants BLAST match Identity % A45 16S gene Pseudomonas fluorescens 97 (AB680178.1) D1 16S gene Pseudomonas fluorescens 96 (AB680178.1) F72 16S gene Pseudomonas fluorescens 99 (AB680178.1) F87 16S gene Pseudomonas fluorescens 95 (AB680178.1) F92 16S gene Pseudomonas fluorescens 97 (AB680178.1)

Table 6: Sequence data showing the BLAST match for the mutant sequences

109

Growth curve comparing the growth rate of wild type vs mutant 0.3

0.25

0.2

0.15

0.1

Optical density nm) density (600 Optical 0.05

0 0 12 24 36 48 60 Time (hrs)

LEw-24 A45 F87 F72 D1 F92

Fig 17. Changes in optical density (Y axis) over time (X-aixs) for the growth curve of LEw-24 and its mutants A45, F87, F72, D1 and F92.

110

111

Fig 18: HPLC/MS figures for the identification of the degradation products and intermediates

112

References

1. Barathi, S and Vasudevan, N (2001). Utilization of petroleum hydrocarbons

by Pseudomonas fluorescens isolated from a petroleum-contaminated soil. Environment

International 26 (5-6):413-416.

2. Bourne, D., Jones, G., Blakeley, R and Jones, A (1996). Enzymatic pathway for the

bacterial degradation of the cyanobacterial cyclic peptide toxin microcystin LR.. American

Society of Microbiology 62 (11):4086-4094.

3. Bourne, D., Riddles, P., Jones, G., Smith, W and Blakeley, R (2001). Characterisation of a

gene cluster involved in bacterial degradation of the cyanobacterial toxin microcystin LR.

Environmental Toxicology 16(6): 523-534.

4. Dietrich, D and Hoeger, S (2005). Guidance values for microcystins in water and

cyanobacterial supplement products (blue-green algal supplements): a reasonable or

misguided approach? Toxicology and Applied Pharmacology 203(3): 273-289.

5. Falconer, I and Yeung, D (1992). Cytoskeletal changes in hepatocytes induced

by Microcystis toxins and their relation to hyperphosphorylation of cell proteins. Chemico-

Biological Interactions 81(1-2):181-196.

6. Fontanillo, M and Kohn, M (2018). MCs: Synthesis and structure–activity relationship

studies toward PP1 and PP2A. Bioorganic and Medicinal Chemistry 26(6):1118-1126.

113

7. Jones, G and Orr, P (1994). Release and degradation of microcystin following algaecide

treatment of a Microcystis aeruginosa bloom in a recreational lake, as determined by HPLC

and protein phosphatase inhibition assay. Water Research 28:871-876.

8. Kormas, K and Lymperopoulou, D (2013). Cyanobacterial toxin degrading bacteria: who

are they? BioMedical Research International 2013 ( 2013): 1-12

9. .Park, H., Sasaki, Y., Maruyama, T., Yanagisawa, E., Hirashi, A., Kato, K (2001).

Degradation of the cyanobacterial hepatotoxin microcystin by a new bacterium

isolated from a hypertrophic lake. Environmental Toxicology 1(1):337-344.

10. Manage P, Edwards C, Singh B, Lawton L (2009) Isolation and identification of novel

microcystin-degrading bacteria. Appl Environ Microbiol 75:6924–6928.

11. Mou X., Lu, X., Jacob, J and Heath, R (2013). Metagenomic identification of

bacterioplankton taxa and pathways involved in microcystin degradation in Lake Erie.

PLoS ONE 8(4):1-14.

12. Petti, C., Polage, C and Schreckenberger, P (2005). The Role of 16S rRNA Gene

Sequencing in Identification of Microorganisms Misidentified by Conventional Methods.

Journal of Clinical Microbiology 43(12):6123-6125.

13. Peregrin-Alvarez, J., Sanford, C and Parkinson, J (2009). The conservation and

evolutionary modularity of metabolism. Genome Biology 10(63): 1-17.

14. Puddick, J., Prinsep, M., Wood, S., Cary, S., Hamilton, D and Holland, P (2015). Further

114

Characterization of Glycine-Containing Microcystins from the McMurdo Dry Valleys of

Antarctica. Toxins 7(2):493-515.

15. Sangolkar, L., Maske, S and Chakrabarti, T (2006). Methods for determining

microcystins (peptide hepatotoxins) and microcystin-producing cyanobacteria. Water

Research 40(19):3485-3496.

16. Shamsollahi, H., Alimohammadi, M., Nabizadeh, R ., Nazmara, S and Mahvi, A

(2015). Measurement of Microcystin -LR in Water Samples Using Improved HPLC

Method. Global Journal of Health Science 7(2): 66-70.

17. Somdee, T., Thunders, M., Rucks, M., Allison, M and Page R (2013) Degradation of [Dha

(7)] MC-LR by a microcystin degrading bacterium isolated from Lake Rotoiti, New

Zealand. ISRN Microbiology 2013:1-8.

18. Yang, F., Zhou, Y., Yin, L., Zhu, G., Liang, G and Pu, Y (2014). Microcystin-Degrading

Activity of an Indigenous Bacterial Strain Stenotrophomonas acidaminiphila MC-LTH2

Isolated from Lake Taihu. PLOS One 9(1): 1-7.

115

Chapter V: Identification and characterization of microcystin induced apoptosis in HepG2 cells

Preface

The fifth chapter for my dissertation has been made into a manuscript for publication and has been submitted to Toxicon. My co-author, Gary Koski has helped me greatly in terms of the experiment set up, data interpretation and culturing of the cells. My dissertation advisor, Xiaozhen

Mou has advised me a lot with the data and writing of my paper.

Abstract

Microcystins (MCs) are a class of hepatotoxins that are commonly produced by freshwater cyanobacteria. MCs harm liver cells through inhibiting protein phosphatases 1 and 2A (PP1 and

PP2A) and can produce dualistic effects, i.e., apoptosis and uncontrolled cellular proliferation. The induction of apoptosis in MC treated hepatic cells has been described previously; however, the apoptotic pathway promoted by MC remains unclear. To address this, HepG2 human hepatoma cells were exposed to MC-LR, the most prevalent isomer of MCs, morphological and physiological responses were examined. Microscopy and Alamar Blue assay showed that HepG2 cells responded to MC-LR treatment with apoptosis characteristics, such as clumping and shrinking of cells and detachment from the culture surface. A fluorescent caspase activation assay further revealed activation of all tested apoptosis-dependent caspases (i.e., caspase-3/7, 8 and 9) after 24 hrs of 116

MC-LR treatment. Furthermore, caspase-8 was found being activated 4 hrs after MC-LR treatment, earlier than observed activation of caspase-9 (8 hrs after MC-LR treatment). These data suggest that MC-LR can induce apoptosis of HepG2 cells through both extrinsic and intrinsic pathways and that the extrinsic pathway may be activated before the intrinsic pathway. This knowledge can be potentially used for developing treatments for MC exposed hepatic cells and to develop a better understanding of MC cytotoxicity.

Key words: Microcystin, HepG2, apoptosis, caspase

Introduction

Extensive growths, or blooms, of cyanobacteria in freshwater environments are hazardous to ecosystem functions and human health (Paerl et al., 2013). One major harmful effect of cyanobacterial blooms is the production of secondary metabolites, or cyanotoxins (Blaha et al.,

2009). The most prevalent cyanotoxins in freshwater systems are microcystins (MCs), a group of potent liver toxins. MCs are produced by many common freshwater cyanobacteria, such as

Microcystis, Anabaena and Planktothrix (Hisbergues et al., 2003). More than 90 MC isoforms have been identified and all possess a cyclic heptapeptide structure, which consists of five constant and two variable amino acid residues (Niedermeyer et al., 2013). The most abundant MC form is

MC-LR, which has leucine (L) and arginine (R) as variable amino acid residues (Paerl et al., 2013).

The unique cyclic structure of MCs makes them resistant to physical and chemical breakdown in 117

natural environments (Chen et al., 2016) and their removal from freshwaters is highly dependent on microbial activities (Mou et al., 2013; Krishnan et al., 2018).

Humans can be exposed to MCs via multiple routes, including ingestion of, aerosol inhalation of and dermal contact with toxin-contaminated water as well as consumption of foods that have been irrigated by (such as vegetables) or lived in (such as fish and shellfish) contaminated waters (Watanabe et al., 1988; Magalhaes et al., 2015). Once entering the human body, MCs bind to organic anion transporter polypeptides (OATPs), which inhibits activities of protein phosphatases 1 and 2a (PP1 and PP2a; Campos et al., 2010). OATPs are abundant on liver cells; this makes liver a primary susceptible organ for MC toxicity (Azevedo et al., 2002). Chronic exposes to low concentrations of MCs in drinking and recreational water have been identified as a risk factor for primary liver cancer (PLC) for humans (Ueno et al., 1996).

Exposure to sudden high dosages of MCs can result in death of eukaryotic cells. A recent study suggests that MC-LR induced liver cell death is through necrosis rather than apoptosis in both of in vivo mice model and in vitro human hepatocytes (Woodbright et al., 2017). However, apoptosis has been suggested as the outcome for MC acute toxicity from many more studies using a variety of cell lines, such as mice liver cells (Weng et al., 2007), Sertoli cells (Li et al., 2012;

Huang et al., 2016) and human kidney cells (Piyathilaka et al., 2015) (Fig 3). Studies have also indicated that MC exposure may stimulate production of reactive oxidative species (ROS), which decrease mitochondrial permeability potential (MMP) and lead to apoptosis via the intrinsic pathway (Weng et al., 2007; Huang et al., 2016; Li et al., 2016) (Fig 3). However, a few studies 118

have also suggested the presence of the extrinsic apoptosis via the NF-κB pathway (Feng et al.,

2011; Ji et al., 2011) (Fig 3).

The lack of consensus for the pathway of MC-induced cell death calls for more studies on this topic to obtain a comprehensive understanding. In this study, acute MC toxicity was found to induce apoptosis in human HepG2 cells based on examinations on cell metabolic activity and morphological changes. Activation of apoptosis-specific caspases was further examined to further elucidate the role of intrinsic and extrinsic pathways in MC induced apoptosis.

Materials and methods

Culturing of HepG2 cells

HepG2 cells were grown in T-75 Falcon culture flasks (Fisher Scientific, Waltham, MA) with Dulbecco's minimum essential medium (DMEM; Thermo Fisher Scientific, Waltham, MA,

USA) that had been supplemented with 10% foetal bovine serum (FBS, Atlanta Biologicals,

Flowery Branch, GA, USA), 1% penicillin/streptomycin (Thermo Fisher Scientific, Waltham, CA,

USA) and 4.5g/L L-glutamine-sodium pyruvate (Gibco, Gaithersburg, MD, USA). The same medium was also used in multiple steps for the rest of the experiments and is referred to as modified DMEM medium. Cells were incubated in the dark at 37ºC and with 5% CO2 for 48-72 hrs until 90% confluence of growth was observed. HepG2 cells were counted using a haematocytometer (Abcam, Cambridge, MA, USA) and then adjusted to 106 cells/ml using the same culturing media for subsequent analysis.

Trypsinization and Alamar Blue Assay 119

Fresh HepG2 cells (25 ml at 106 cells/ml) were incubated with 3 ml of trypsin solution

(1×pancrease derived trypsin in EDTA; Thermo Fisher Scientific, Waltham, MA, USA) and incubated at 37ºC and 5% CO2 for 10 min then mixed with 10 ml of modified DMEM medium

(Rampersad et al., 2012). Afterwards, trypsinized HepG2 cells were centrifuged at 750 ×g for 10 min at 20ºC. The supernatant was discarded; resulting cell pellets were then re-suspended by repeated pipetting in 2 ml of modified DMEM medium. Re-suspended cells were counted using a hemocytometer (Abcam, Cambridge, MA, USA) and adjusted to a concentration of 1×106 cells/ml.

Trypsinized HepG2 cells (1×106 cells/ml; 200 µl/well) were transferred into 96-well microplates (Sigma-Aldrich, St.Louis, MO, USA) and incubated at the same condition as described above for 24 hrs (Rampersad et al., 2012). Afterwards, 30 µl of MC-LR (Cayman

Chemicals, Ann Arbor, MI, USA) was added to each well at final concentrations of 0, 0.1, 1, 5,

10, 15, 25, 30 or 35 µM and incubated for another 24 hrs. At the end of MC-LR treatment, Alamar

Blue reagent (10 µl; Thermo Fisher Scientific, Waltham, MA, USA) was mixed into each well and the plates were incubated again at the same condition for 10 hrs. At the end of incubation, absorbance readings were taken at 600 nm using a Synergy Multimode Reader (Bioteck, Winooski,

VT, USA).

Microscopic observation

Fresh HepG2 cells (25 ml at 106 cells/ml) were amended with MC-LR at a final concentration of 0 (as controls) or 5 µM and incubated at 37ºC in 5% CO2 for 24 hrs. After

120

incubation, cell morphology and internal structures were examined using a Fluoview 300 confocal microscope (Olympus, Pittsburgh, PA, USA).

Flow cytometric analysis for caspase activity

Trypsinized HepG2 cells (25 ml at 106 cells/ml) were prepared as above and then incubated with 5µM MC-LR for 24 hrs in modified DMEM medium at the same conditions described above.

Afterwards, HepG2 cells were further processed using a CellEvent Caspase-3/7 Green Flow

Cytometry Assay Kit (Fisher Scientific, Waltham, MA, USA) following the manufacturer’s protocol. The final staining step involved incubating processed HepG2 cells with diluted green detection reagent at 37ºC and with 5% CO2 for 24 hrs. The stained cells were washed five times using the provided wash buffer and then examined using a FacsAria Flow Cytometer (Becton

Dickinson, San Jose, CA, USA). The fluorescence emitted by cells with active caspase-3 was detected using the FITC channel.

Similar procedures were carried out to examine activities of caspase-8 and caspase-9 using a Vybrant FAM Caspase-8 Assay Kit (Fisher Scientific, Waltham, MA, USA) and a Caspase-9

(active) FITC Staining Kit (Abcam, Cambridge, MA, USA), respectively.

To track potential lags between the expression time of caspase-8 and -9, the flow cytometry assays were repeated following the steps described above using trypsinized HepG2 cells that had been treated with 5µM MC-LR for 0, 4, 8 or 12 hrs.

121

Statistical analysis

Statistical analyses were carried out using R software (R core team, 2013). A T-test was performed comparing the optical density and cell count values between the MC-LR treated samples and controls in both of the Alamar Blue assay and flow cytometry assay. Significant differences were reported when p <0.05.

Results & discussion

In vitro viability after MC-LR treatment

Alamar Blue Assay was used to quantify in vitro viability of HepG2 cells after being exposed to various concentrations of MC-LR. Following a 24-hr incubation with MC-LR, HepG2 cells that received MC-LR at concentrations of 0.1 and 1 µM reduced Alamar Blue significantly

(p<0.05) and at levels similar to control HepG2 cells that received no MC-LR (Fig 19). These suggest that metabolic activity of tested HepG2 cells was not significantly affected by MC-LR at concentrations of 1 µM or lower (Fig 19). In contrast, when concentration of MC-LR increased to

5 µM and higher, treated HepG2 cells showed from minor (5µM; p<0.05) to no reduction (10-35

µM) of Alamar Blue optical density. These suggest that metabolic activities of HepG2 cells were significantly inhibited by MC-LR at 5 µM and higher. Therefore, all the subsequent experiments in this study were performed using MC-LR at 5 µM.

Total concentration of MCs during cyanobacterial harmful algal blooms (CyanoHABs) often reach over 0.1 nM or ~100 µg/L in highly eutrophic freshwater lakes (Fromme et al., 2000;

122

Dumouchelle and Stelzer, 2014). However, through bioaccumulation, intracellular MC concentrations can surge to tens of thousands µg/g dry weight in fishes, invertebrates and zooplankton from CyanoHAB impacted lakes (Magalhaes et al., 2003; Ferrão-Filho and

Kozlowsky-Suzuki, 2011). Therefore, the tested MC-LR is within the concentration range that human consumers can potentially encounter.

Previously, inhibition of cell activity by MC-LR has been primarily been studied in non- liver cells, even though hepatic cells are the primary target for MC-LR (Campos et al., 2010).

These in vitro studies also mostly used MC-LR at 20 µM or higher (Botha et al., 2004; Campos et al., 2010), with lesser number of studies focusing on human liver cells where apoptosis is induced at a low MC-LR concentration (Basu et al., 2018). The suggested higher susceptibility of HepG2 cell line in our study is likely due to the abundance presence of OATP, resulting in apoptosis induction at a lower MC-LR concentration as seen in a recent study (Basu et al., 2018).

MC-LR induced cellular morphology changes

MC-LR treated HepG2 cells showed lower confluency with a high number of rounded cells that had detached from the surface of the culture flask. Confocal microscopic analysis further revealed that MC-LR treated HepG2 cells were shrunken and clumped (Fig 20a). These observed features matched typical characteristics of cells entering apoptosis stage (Cummings et al., 2013).

A recent study has suggested that MC-LR cause liver cell death through necrosis (Woodbright et al., 2017). However, typical morphological changes associated with necrosis are swelling of the organelles accompanied by formation of vacuoles (Ziegler et al., 2004), which weren’t observed 123

in MC-LR treated hepatic cells in our study. Control HepG2 cells, on the other hand, displayed a healthy growth with a confluent monolayer (Fig 20b). Therefore, our results agreed with the common consensus that acute exposure of MC-LR can lead to cell apoptosis.

Caspase expression in response to MC-LR exposure

Activity of caspase-3 and -7, executioner caspases that are shared by both intrinsic and extrinsic apoptosis (Fig. 3; Elmore et al., 2007), was examined in MC-LR treated HepG2 cells using the CellEvent Caspase3/7 green flow cytometry (FCM) assay. Activation of targeted caspases would produce bright fluorogenic signals in tested cells. FCM histogram analysis illustrated clear upshifts of fluorescence signal of MC-LR treated HepG2 cells from untreated cells

(Fig 21a; p < 0.05), indicating significant up-regulation of caspase-3/7 activity. With slight differences, both capspase-3 and -7 induce DNA fragmentation, protein cross-link, cell dysmorphology and other biochemical events that are required for cell demolition and completion of apoptosis (Porter et al., 1999; Walsh et al., 2008). Therefore, results of caspase-3/7 FCM analysis were in line with those of Alamar Blue assay and microscopy analysis of cell morphology change and consistently suggest that MC-LR carry apoptosis-dependent cytotoxicity to human liver HepG2 cells. This phenomenon has previously only been identified in primary hepatocytes

(Lone et al., 2016) and some permanent cell lines (Zhou et al., 2017).

Extrinsic and intrinsic apoptosis pathways differ in types of initiator caspases (Fig 3). The extrinsic pathway is initiated by caspase-8, which is activated by death receptors (Ghobrial et al.,

2015). In contrast, the intrinsic pathway is initiated by caspase-9, which is activated by 124

mitochondria-mediated processes (Scoltock et al., 2004; Fulda et al., 2006). To specify which pathway MC-induced apoptosis would take, activities of above two caspases were examined separately using flow cytometric analyses (FCM). Compared with their corresponding controls,

MC-treated HepG2 cells illustrated much higher fluorescence intensities in FCM analyses for both of caspase-8 (Fig. 21b) and -9 (Fig. 21c), illustrating both intrinsic and extrinsic pathways of apoptosis were triggered by MC-LR in HepG2 cells. Previous studies have examined and identified both of intrinsic and extrinsic pathways separately in various MC-LR treated cell lines

(Feng et al., 2011; Ji et al., 2011; Zhang et al., 2013; Huang et al., 2016). Our results, however, served as one of the first evidence to show both pathways are simultaneously induced in human liver cells by acute MC exposure.

Induction of both intrinsic and extrinsic apoptotic pathways has been observed before for other types of toxin factors, for example Bifidobacterim infantis (Yin et al., 2013) and exotoxin producing uropathogenic E.coli (Klumpp et al., 2006). In these cases, extrinsic pathway was directly induced and the intrinsic pathway is then indirectly activated via the BID cleavage process

(Klumpp et al., 2006). However, other apoptosis studies have suggested that when both intrinsic and extrinsic pathways are involved, intrinsic pathway may be activated independently (Lamkanfi and Dixit, 2010).

To further examine this aspect, a time series sampling and FCM analysis were further performed to examine potential lag time between activation of intrinsic and extrinsic pathways

(Fig. 22). The FCM histogram demonstrated that activation of caspase-8 was observed as early as 125

4 hrs following MC-LR treatment (Fig. 22a). In contrast, activation of caspase-9 was observed in

HepG2 cells at a later sampling point (8 hrs; Fig.22b) of MC-LR treatment. Once activated, both caspase-8 and-9 maintained at similar high levels until the end of the experiment. The observed sequential activation of capsase-8 and then caspase-9 suggest that extrinsic pathway is activated upstream of intrinsic pathway, supporting the hypothesis that MC-LR induced intrinsic apoptosis is activated by extrinsic pathway (Zhao et al., 2010). However, we cannot entirely rule out the possibility that extrinsic and intrinsic pathways are stimulated by MC-LR independently

(Elankumaran et al., 2006) and extrinsic pathway is more sensitive than intrinsic pathway for MC-

LR induced apoptosis.

Conclusion

In conclusion, this study demonstrated that MC-LR could induce a complex apoptosis in liver cells that combines both intrinsic and extrinsic pathways. Extrinsic pathway, i.e., caspase-8, was activated earlier than intrinsic pathway, and could potentially serve as target to develop therapeutic treatment for MC-LR intoxication.

Acknowledgement

We would like to thank Drs. Preeti Pathak and John Chiang from North East Ohio Medical

University (NEOMED) for providing us with the HepG2 cells required for the experiments. This project was supported by Kent State University and National Sciences Foundation (Award number

1605161).

126

0.9

) 0.8

0.7

0.6 0.5 0 hr

0.4 10 hr 0.3

nm (630 density Optical 0.2 0.1

0 0 0.1 1 5 10 15 25 30 35 Microcystin concentration (µM)

Fig 19: Effects of various concentrations of MC-LR on metabolic activity of HepG2 cells with optical density values before (0 hr) and after (10 hr) incubation with Alamar Blue.

127

10µm

)

10µm

10µ

m Fig 20: Confocal microscopy pictures of control (untreated) (a) and 5µM MC-LR treated (b) HpeG2 cells after 24 hrs of incubation. 128

a) Caspases 3/7 b) Caspases 8 c) Caspases 9

Cell number Cell

FITC –A (Fluorescent intensity)

Fig 21: Histogram showing fluorescence for untreated HepG2 cells (black) grown in DMEM medium and MC-LR treated (blue) HepG2 cells stained with caspases 3/7 specific binding dye (green) (a), caspases 8 specific stain (red) (b) and caspases 9 FITC stain (blue) (c) . .

129

a) Caspase-8

8 hr 12 hr 0 hr 4 hr

Cell number Cell

b) Caspase-9

4 hr 8 hr 12 hr 0 hr

FITC –A (Fluorescent intensity)

Fig 22: Histogram representing the fluorescence for treated, caspase-8 (red, a) and caspase-9 expressing (blue, b) HepG2 cells, observed at different time points of 0, 4, 8, 12 and 24 hrs of MC-LR treatment. Control (black, untreated) HepG2 cells are also shown in this figure.

130

References

1. Azevedo, S., Carmichael, W., Jochimsen, E., Rinehart, K., Lau, S., Shaw, G and

Eaglesham, G., 2002. Human intoxication by microcystins during renal dialysis treatment

in Caruaru—Brazil. Toxicology. 181-182:441-446.

2. Basu, A., Dydowiczová, A., Čtveráčková, L., Jaša, L., Trosko, J., Bláha, L and

Babica, P., 2018. Assessment of hepatotoxic potential of cyanobacterial toxins using 3D

In Vitro model of adult human liver stem cells. Environmental Science &

Technology. 52(17):10078-10088.

3. Blaha, L., Babica, P and Marsalek, B., 2009. Toxins produced in cyanobacterial water

blooms – toxicity and risks. Interdisciplinary Toxicology. 2(2):36-41.

4. Botha, N., Gehringer, M., Downing, T., Van de Venter, M., Shephard, E., 2004. The role

of microcystin-LR in the induction of apoptosis and oxidative stress in CaCo2 cells.

Toxicon. 43(1):85-92.

5. Campos, A and Vasconcelos, V., 2010. Molecular mechanisms of microcystin toxicity in

animal cells. 11(1):268-287.

6. Chen, L., and Xie, P., 2016. Mechanisms of microcystin-induced cytotoxicity and

apoptosis. Mini-Reviews in Medicinal Chemistry. 16 (13):1018-1031.

131

7. Cummings, B., Wills, L and Schnellmann, R., 2013. Measurement of cell death in

mammalian cells. Current Protocol of Pharamacology. 1(12):1-30.

8. Dumouchelle, D and Stelzer, E., 2014. Chemical and biological quality of water in Grand

Lake St. Marys, Ohio, 2011–2012, with emphasis on cyanobacteria. U.S. Geological

Survey Scientific Investigations Report 2014. 5210(2014):51-60.

9. Elankumaran, S., Rockemann, D and Kamal, S., 2006. Newcastle disease virus exerts

oncolysis by both intrinsic and extrinsic caspase dependent pathways of cell death. Journal

of Virology. 80(15):7522-7534.

10. Elmore, S., 2007. Apoptosis: A review of programmed cell death. Toxicology Pathology.

35(4): 495-516.

11. Feng, G., Abdalla, M., Li, Y., Bai, Y., 2011. NF-κB mediates the induction of Fas

receptor and Fas ligand by microcystin-LR in HepG2 cell. Molecular and Cellular

Biochemistry. 352(1-2): 209-219.

12. Ferrão-Filho, S and Kozlowsky-Suzuki, B., 2011. Cyanotoxins: bioaccumulation and

effects on aquatic animals. Marine Drugs. 9(12):2729-2772.

13. Fromme, H., Köhler, A., Krause, R. and Führling, D., 2000. Occurrence of cyanobacterial

toxins—microcystins and anatoxin a—in Berlin water bodies with implications to human

health and regulations. Environmental Toxicology: An International Journal. 15(2):120-

130.

132

14. Fulda, S and Debatin, K., 2006. Extrinsic versus intrinsic apoptosis pathways in anticancer

chemotherapy. Oncogene 25 (34):4798-4811.

15. Ghobrial, M.G., H.S. Nassr, and A.W. Kamil., 2015. Bioactivity effect of two macrophyte

extracts on growth performance of two bloom-forming cyanophytes. The Egyptian Journal

of Aquatic Research. 41(2015):69-81.

16. Hisbergues, M., Christiansen, G., Rouhiainen, L., Sivonen, K and Börner, T., 2003. PCR-

based identification of microcystin-producing genotypes of different cyanobacterial

genera. Archives of Microbiology. 180 (6):402-410.

17. Huang, H., Liu, C., Fu, X., Zhang, S., Xin, Y., Li, Y., Xue, L., Cheng, X and Zhang, H.,

2016. Microcystin-LR induced apoptosis in rat Sertoli cells via the mitochondrial

caspase-dependent pathway: Role of reactive oxygen species. Frontiers in Physiology.

7(397):1-10.

18. Ji, Y., Lu, G., Chen, G., Huang, B., Zhang, X., Shen, K and Wu, S., 2011. Microcystin-

LR Induces Apoptosis via NF-κB /iNOS Pathway in INS-1 Cells. International Journal of

Molecular Science. 12(7):4722-4734.

19. Klumpp, D., Rycyck, M., Chen, M., Thumbikat, P., Sengupta, S and Schaeffer, A., 2006.

Uropathogenic Escherichia coli induces extrinsic and intrinsic cascades to initiate

urothelial apoptosis. Infection and Immunity. 74(9):5106-5113.

133

20. Krishnan, A., Zhang, Y and Mou, X., 2018. Isolation and characterization of microcystin-

degrading bacteria from Lake Erie. Bulletin of Environmental Contamination and

Toxicology. 101(5):617-623.

21. Lamkanfi, M and Dixit, V., 2010. Manipulation of Host Cell Death Pathways during

Microbial Infections. Cell Host & Microbe. 8(1): 44-54.

22. Li, Y and Han, X., 2012. Microcystin–LR causes cytotoxicity effects in rat testicular Sertoli

cells. Environmental Toxicology and Pharmacology. 33(2):318-326.

23. Li, Y., Li, J., Huang, H., Yang, M., Zhuang, D., Cheng, X., Zhang, H and Fu, X., 2016.

Microcystin-LR induces mitochondria-mediated apoptosis in human bronchial epithelial

cells. Experimental and Therapeutic Medicine. 12(2):633-640.

24. Lone, Y., M. Bhide, and R.K. Koiri., 2016. Microcystin-LR induced immunotoxicity in

mammals. Journal of Toxicology. 2016 (2016):1-5.

25. Magalhaes, V., Soares, R and Azevedo, S., 2001. Microcystin contamination in fish from

the Jacarepaguá Lagoon (Rio de Janeiro, Brazil): ecological implication and human health

risk. Toxicon 7(1):1077-1086

26. Magalhaes, V., Marinho, M., Domingos, P., Oliveira, A., Costa, S., Azevedo, L and

Azevedo, S., 2003. Microcystins (cyanobacteria hepatotoxins) bioaccumulation in fish and

crustaceans from Sepetiba Bay (Brasil, RJ). Toxicon, 42(3): .289-295.

134

27. Mou X, Lu X, Jacob J, Heath R., 2013b. Metagenomic identification of bacterioplankton

taxa and pathways involved in microcystin degradation in Lake Erie. PLoS ONE. 8(4):1-

14.

28. Niedermeyer, T., 2013. Microcystin congeners described in the literature. Available online:

http://dx.doi.org/10.6084/m9.figshare.880756.

29. Paerl, W., Otten, G., 2013.Harmful cyanobacterial blooms: causes, consequences and

controls. Microbial Ecology 65(4):995-1010.

30. Piyathika, M., Pathmalal, M., Tennekoon, H., DeSliva, B., Samarakoon, and Chanthirika,

S., 2015. Microcystin-LR-induced cytotoxicity and apoptosis in human embryonic kidney

and human kidney adenocarcinoma cell lines. Microbiology. 161 (4):819-828.

31. Porter, A and Janicke, R., 1999. Emerging roles of caspase-3 in apoptosis. Cell Death and

Differentiation. 6(2):99-104.

32. Rampersad, S., 2012. Multiple applications of Alamar Blue as an indicator of metabolic

function and cellular health in cell viability bioassays. Sensors 12(9): 12347-12360.

33. Scoltock, A.B., and J.A. Cidlowski., 2004. Activation of intrinsic and extrinsic pathways

in apoptotic signaling during UV-C-induced death of Jurkat cells: the role of caspase

inhibition. Experimental Cell Research. 297(1):212-223.

34. Ueno Y., S. Nagata, T. Tsutsumi, A. Hasegawa, M.F. Watanabe, H.D. Park, G.C. Chen, G.

Chen, and SZ. Yu., 1996. Detection of microcystins, a blue-green algal hepatotoxin, in

135

drinking water sampled in Haimen and Fusui, endemic areas of primary liver cancer in

China, by highly sensitive immunoassay. Carcinogenesis. 17(6):1317-1321.

35. Watanabe, M., S. Oishi, K. Harada, K.Matsuura, H. Kawai, and M. Suzuki., 1988. Toxins

contained in Microcystis species of cyanobacteria (blue-green algae). Toxicon. 26

(11):1017-1025.

36. Weng, D., Y. Lu, Y. Wei, Y. Liu, and P. Shen., 2007. The role of ROS in microcystin-LR-

induced hepatocyte apoptosis and liver injury in mice. Toxicology. 232(1-2):15-23.

37. Woolbright, B., Williams, C., Ni, H., Kumer, S., Schmitt, T., Kane, B and Jaeschke, H.,

2017. Microcystin-LR induced liver injury in mice and in primary human hepatocytes is

caused by oncotic necrosis. Toxicon. 125(2017):99-109.

38. Yin, X., Yu, B., Tang, Z., He, B., Ren, J., Xiao, X and Tang, W., 2013. Bifidobacterium

infantis-mediated HSV-TK/GCV suicide gene therapy induces both extrinsic and intrinsic

apoptosis in a rat model of bladder cancer. Cancer Gene Therapy. 20:77-81.

39. Ziegler, U and Groscurth, P., 2004. Morphological features of cell death. News in

Physiological Society. 19(3):124-128.

40. Zhao, Y., Li, R., Xia, W., Neuzli, J., Lu, Y., Zhang, H., Zhao, X., Zhang, X., Sun, C and

Wu, K., 2010. Bid integrates intrinsic and extrinsic signaling in apoptosis induced by α-

tocopheryl succinate in human gastric carcinoma cells. Cancer Letters. 288(1):42-49.

41. Zhou, Y., Xu, X., Yu, B AND Yu, G., 2017. Characterization of in vitro effects of

microcystin-LR on intestinal epithelial cells. Environmental Toxicology. 32(5):1539-1547 136

GENERAL DISCUSSION

This dissertation aimed to develop a better understanding of the biological degradation mechanisms and toxicological effects of MC-LR. With increasing number of freshwater lake resources suffering from the issues of cyanobacteria blooms (Smith et al., 1999) and microcystin contamination problem become more prevalent, it is critical that we understand the role of bacteria in biological eradication of microcystins (Lawton et al., 2011). My dissertation study examined the taxonomic and functional diversity of bacteria with microcystin degradation potentials from

Lake Erie (Chapters 2 & 3) and the pathway governing this process (Chapter 4). We found that microcystin degrading bacteria from Lake Erie belonged to proteobacteria, actinobacteria and firmicutes phylum. MC- degrading bacteria from Lake Erie adopt a unique degradation pathway as opposed to the previously identified mlr degradation pathway (Mou et al., 2013). In Chapter 4, potential pathway for MC degradation via non-mlr genes was investigated by using transposon mutagenesis and HPLC/MS. We found that bacterioplankton from Lake Erie adopt a novel microcystin degradation pathway governed by genes lying adjacent to the 16S rRNA region of the bacteria. We also identified the two potential degradation products of MC-LR seen using

HPLC/MS. Identification of the apoptotic effects of microcystin in mammalian cell culture has been a topic of extreme interest due to the toxicity of MC-LR (Campos et al., 2010; Grabow et al.,

1982). In this dissertation, we have been able to successfully identify the morphological changes accompanying exposure of HepG2 cells to microcystin (Chapter 5). We were also able to identify 137

the involvement of both extrinsic and intrinsic caspases dependent pathway in microcystin induced apoptosis in HepG2 cells. The subsequent sections will provide a synthesized review of results obtained for each subject and their significance.

General characterization of culturable microcystin degrading bacteria in Lake Erie

In this study we confirmed the presence of multiple microcystin degrading bacteria from the surface water of Lake Erie. They belong to diverse taxa within the phylum of proteobacteria, actinobacteria and firmicutes with great diversity in phenotypic characteristics of color, morphology, motility and biofilm formation. It was also observed that the degradation rate of these bacteria were affected by abiotic factors of pH and temperature (Chapter 2 and 3). However, it was observed that addition of organic nutrient did not change the rate of microcystin degradation in bacteria from Lake Erie except for isolate LEw-2029 which shows an increase in microcystin degradation rate. Biotic factors in the form of interspecific interactions between two different MC degrading bacteria isolates were found to increase the rate of microcystin degradation for some isolate combinations. Amplification of mlrA gene via established PCR program showed absence of the mlrA gene in 40 MC-degrading bacteria from Lake Erie. These results suggest the absence of the mlrA gene for MC-degrading bacteria from Lake Erie. Similar results looking at the absence of mlrA gene was observed for other microcystin degrading bacteria (Yang et al., 2014; Mou et al., 2013; Tsao et al., 2017). 138

Effects of pH variation was also observed with highest microcystin degradation rate recorded at pH 7-8, with some isolate showing degradation at pH 9. Freshwater lake resources affected by blooms typically are typically have pH values ranging between 9-10 (Paerl et al.,

2011), which makes it an ideal environment for functional bacteria to degrade microcystin.

Degradation by our isolates was also found to be at faster rates at the highest testing temperatures, i.e., 35ºC which is typically the temperature of the surface water during a cyanobacterial bloom in summer. These results help us identify the optimal pH, temperature and organic nutrient concentration, under which optimal microcystin degradation is observed. Identifying the optimal degradation conditions for the MC-degrading bacteria, make them ideal candidates for biological degradation in Water Treatment Plant (WTP) However, further work will be needed, using microcystin contaminated water from lakes to study the efficiency of these bacteria when exposed to natural waters.

Non-mlr pathway for Microcystin degradation

With this dissertation, I was successfully able to identify the presence of alternate microcystin degradation genes and intermediates in MC-degrading bacteria from Lake Erie. A novel microcystin degradation pathway was screened in MC-degrading bacteria present in Lake

Erie. The genes responsible for degradation were found to lie adjacent to the 16S rRNA region of the bacteria. Two degradation intermediates A & B were identified for the MC-LR degradation process. Degradation products A and B were identified as having m/z ratio of 509.2 and 1061

139

respectively. Degradation product B was found to be present in the suspension after complete removal of the MC-LR within 10 hrs of incubation.

Confirmatory tests for presence of microcystin degradation in wild type and its absence in mutants were also performed to ensure the mutation of microcystin degrading gene. These results supports other studies identifying the absence of mlrA degrading gene and the presence of other degradation genes present in bacteria from Lake Erie (Yang et al., 2014; Mou et al., 2013).

Identification of this novel microcystin degradation pathway present in MC-degrading bacteria from Lake Erie (Chapter 4) will enable us to get a better understanding of the mechanism involved in the microcystin degradation performed by bacteria from Lake Erie. The genes responsible for microcystin degradation can be mutated and the degradation process enhanced.

An unexpected finding in this study was the identification of microcystin degradation gene being present in the 16S rRNA region of the MC-degrading bacteria. This region has been attributed to only taxonomic identification of bacteria and shows presence of no other expression of protein (Janda et al., 2007). But, in the current study we observed the presence of microcystin degradation gene being present in the 16S rRNA region of the bacteria. It would be interesting to perform further study on the microcystin degrading gene present in the 16S rRNA gene. This will further elucidate the mechanism underlying microcystin degradation in bacteria.

Hepatotoxic effects of microcystin

In this study we have closely looked at the cytotoxicity of MC-LR in HepG2 hepatoma cells. The morphological changes recorded for MC-LR treated HepG2 cells showed rounding of 140

cells followed by their clumping and shrinking. Alamar Blue assays performed to identify the reduction in metabolic activity for the MC-LR exposed hepatic cells also indicated towards cell death after administration of as little as 5µM of MC-LR after 24 hrs. These changes indicate the induction of apoptosis in MC-LR treated hepatic cells (Saraste et al., 2000). The above observed results are in accordance with the previously seen images for MC-LR exposed hepatic cells

(Campos et al., 2010). The changes in morphology and reduction in metabolic activity after administration of 5µM of MC-LR provide further evidence for triggering of apoptosis after exposure to MC-LR. Induction of apoptosis vs that of necrosis has long been a matter of dispute for MC-LR treated cells (Woolbright et al., 2016). In this study we have performed several assays looking at the presence of caspase after MC-LR exposure, which would help clarify the mechanism of apoptosis adopted by HepG2 cells following MC-LR exposure. The presence of caspase-3, -8 and -9 were recorded using flow cytometric assays along with the time dependent assay showing the activation of caspase-8 before caspase-9. The above results for caspases help conclude that apoptosis induction in MC-LR treated cells occurs via both extrinsic and intrinsic apoptotic pathway. Previously done work have identified the presence of caspase-9 and -3 in MC-LR treated hepatic cells (Gehringer et al., 2003). In this study we can clearly observed the presence of both caspase-8 and -9 in HepG2 cells exposed to MC-LR with the induction of caspase 8 before 9

(Chapter 5).

This unexpected finding of induction of both extrinsic and intrinsic apoptotic pathway in

MC-LR treated hepatic cells help explain the apoptotic phenomenon better. With the help of my 141

findings we will now be able to better understand the mechanism underlying apoptosis induced by

MC-LR in hepatic cells and the overall complexity of its cytotoxicity.

Conclusions and Implications

Microcystin has been recognized as one of the most lethal contaminants in freshwater lake resources (Dawson et al., 1998). This dissertation documented all the findings that are critical for gaining a better understanding of the biodegradation and toxicological effects of MC-LR.

Identifying the different types of bacteria which possess the ability to degrade MC-LR, along with studying the genes and degradation intermediates will help provide a greater understanding of the

MC-LR degradation mechanism. This data will also help recognize the vital role played by these bacteria in complete removal of microcystin from freshwater lake resources. These MC-degrading bacteria can also be utilized for biological degradation of MC-LR in Water Treatment Plants

(WTPs). The successful identification of the apoptotic pathway opens up several opportunities for further exploring the cytotoxicity induced by MC-LR in hepatic cells and also enable in treating liver cells exposed to microcystin.

142

References

1. Campos, A and Vasconcelos, V (2010). Molecular Mechanisms of Microcystin Toxicity

in Animal Cells. International Journal of Molecular Sciences 11(1): 268-287.

2. Dawson, R (1998). The Toxicology of Microcystins. Toxicon 36(7):953-962.

3. Gehringer, M (2003). Molecular Mechanisms of Microcystin Toxicity in Animal Cells.

FEBS Letters 557(1-3): 1-8.

4. Grabow, W., Randt, W., Prozesky, O and Scott, W (1982). Microcystis aeruginosa toxin:

cell culture toxicity, hemolysis and mutagenicity assays. Applied Environmental

Microbiology 43(6): 1425-1433.

5. Janda, J and Abbott, S (2007). 16S rRNA Gene Sequencing for Bacterial Identification in

the Diagnostic Laboratory: Pluses, Perils, and Pitfalls. Journal of Clinical Microbiology

1(1):2761-2764.

6. Lawton, A., Welgamage, A., Manage, M and Edwards, C (2011). Novel bacterial strains

for the removal of microcystins from drinking water. Water Science and Technology 63

(6): 1137-1142.

7. Mou, X., Lu, X., Jacob, J., Sun, S and Heath, R (2013). Metagenomic Identification of

Bacterioplankton Taxa and Pathways Involved in Microcystin Degradation in Lake Erie.

PLOS ONE 8(4):1-14.

143

8. Paerl, H., Hall, N and Calandrino, E (2011). Controlling harmful cyanobacterial blooms

in a world experiencing anthropogenic and climatic-induced change. Science of the Total

Environment 409 (10):1739-1745.

9. Sarsate, A and Pulkki, K (2000). Morphologic and biochemical hallmarks of apoptosis.

Cardiovascular Research 45 (3): 528-537.

10. Smith, V., Tilman, G and Nekola, J (1999). Eutrophication: impacts of excess nutrient

inputs on freshwater, marine, and terrestrial ecosystems. Environmental Pollution 100 (1-

3): 179-196.

11. Tsao, S., Wei, D., Chang, Y and Lee, J (2017). Aerobic biodegradation of microcystin-

LR by an indigenous bacterial mixed culture isolated in Taiwan. International

Biodeterioration & Biodegradation 124:101-108.

12. Nishiwaki-Matsushima, R., Ohta, T., Nishiwaki, S., Suganuma, M and Kohayama, K

(1992). Liver tumor promotion by the cyanobacterial cyclic peptide toxin microcystin-

LR. Journal of Cancer Research and Oncology 118(6): 420-424.

13. Woolbright, B., Williams, C., Ni, H., Kumer, S., Schmitt, T., Kane, B., Jaeschke, H

(2017). Microcystin-LR induced liver injury in mice and in primary human hepatocytes is

caused by oncotic necrosis. Toxicon 125: 99-109.

14. Yang, F., Zhou, Y., Yin, L., Zhu, G and Pu, Y (2014). Microcystin-Degrading Activity of

an Indigenous Bacterial Strain Stenotrophomonas acidaminiphila MC-LTH2 Isolated

from Lake Taihu. PLOS ONE 9(1):1-7. 144

Appendix

Supplementary Table 7: Pure culture isolates from samples collected from a biofilter in Toledo, Akron (water and sediment), Ravenna (water and sludge), Alliance WTP (backwash, sludge) and Sandusky (water) during the summer of 2015 and 2016. Isolates with bold names have the ability to degrade microcystin-LR. TBF 24 Toledo Summer 2016 Bio Filter TBF 23 Toledo Summer 2016 Bio Filter TBF 15B Toledo Summer 2016 Bio Filter TBF 25 Toledo Summer 2016 Bio Filter TBF 26 Toledo Summer 2016 Bio Filter TBF 1113 Toledo Summer 2016 Bio Filter TBF 31 Toledo Summer 2016 Bio Filter TBF 28 Toledo Summer 2016 Bio Filter TBF 18A Toledo Summer 2016 Bio Filter TBF 24 Toledo Summer 2016 Bio Filter TBF 32 Toledo Summer 2016 Bio Filter TBF 19B Toledo Summer 2016 Bio Filter TBF 19A Toledo Summer 2016 Bio Filter TBF64 Toledo Summer 2016 Bio Filter TBF65 Toledo Summer 2016 Bio Filter TBF 19B Toledo Summer 2016 Bio Filter TBF 30 Toledo Summer 2016 Bio Filter TBF 33 Toledo Summer 2016 Bio Filter TBF 34 Toledo Summer 2016 Bio Filter TBF 29 Toledo Summer 2016 Bio Filter TBF 2A Toledo Summer 2016 Bio Filter TBF 2B Toledo Summer 2016 Bio Filter TBF 49 Toledo Summer 2016 Bio Filter TBF 8 Toledo Summer 2016 Bio Filter TBF 21B Toledo Summer 2016 Bio Filter TBF 21A Toledo Summer 2016 Bio Filter TBF 9A Toledo Summer 2016 Bio Filter TBF 20B Toledo Summer 2016 Bio Filter TBF 20A Toledo Summer 2016 Bio Filter

145

S-16-58 Sandusky Summer 2016 Bio Filter 5-16-60 Sandusky Summer 2016 Surface Water 5-16-61 Sandusky Summer 2016 Surface Water S-2127 Sandusky Summer 2016 Surface Water CGB1 Akron 2015 Sediment Akron Water 7 Akron 2015 Water Akron Sediment 1 Akron 2015 Sediment Akron Sediment 2 Akron 2015 Sediment Akron Sediment 3 Akron 2015 Sediment Akron Sediment 4 Akron 2015 Sediment Akron Sediment 5 Akron 2015 Sediment Akron Sediment 6 Akron 2015 Sediment Bouy 1 Akron 2015 Water Bouy 2 Akron 2015 Water Bouy 3 Akron 2015 Water Bouy 4 Akron 2015 Water Bouy 5 Akron 2015 Water Bouy 6 Akron 2015 Water Bouy 7 Akron 2015 Water Bouy 8 Akron 2015 Water Bouy 9 Akron 2015 Water Bouy 10 Akron 2015 Water Bouy 11 Akron 2015 Water Bouy 12 Akron 2015 Water A1 Alliance 2016 Water A2 Alliance 2016 Water A3 Alliance 2016 Water A4 Alliance 2016 Water A5 Alliance 2016 Water A6 Alliance 2016 Water A7 Alliance 2016 Water A8 Alliance 2016 Water A9 Alliance 2016 Water A10 Alliance 2016 Water A11 Alliance 2016 Water 146

A12 Alliance 2016 Water RS1 Ravenna Sludge 2016 Sludge RS2 Ravenna Sludge 2016 Sludge RS3 Ravenna Sludge 2016 Sludge RS4 Ravenna Sludge 2016 Sludge RS5 Ravenna Sludge 2016 Backwash Water ABW 1 Alliance Backwash 2016 Backwash Water ABW 2 Alliance Backwash 2016 Backwash Water ABW 3 Alliance Backwash 2016 Backwash Water ABW 5 Alliance Backwash 2016 Backwash Water AK1 Akron 2016 Water AK2 Akron 2016 Water AK3 Akron 2016 Water AK4 Akron 2016 Water AK5 Akron 2016 Water AK6 Akron 2016 Water AK7 Akron 2016 Water AK8 Akron 2016 Water AK9 Akron 2016 Water AK10 Akron 2016 Water AK12 Akron 2016 Water MC+ W/ON Maumee Bay 2016 Water EMB 6/18 Maumee Bay 2016 Water EMB 6/18 Maumee Bay 2016 Water CGB 9/16 Maumee Bay 2016 Water CGB 9/16 Maumee Bay 2016 Water RSW1 Ravenna Source Water 2016 Water RS5 Ravenna Source Water 2016 Water RS25 Ravenna Source Water 2016 Water RS24 Ravenna Source Water 2016 Water RS23 Ravenna Source Water 2016 Water RS22 Ravenna Source Water 2016 Water RS27 Ravenna Source Water 2016 Water RS26 Ravenna Source Water 2016 Water RL27 Ravenna Sludge 2016 Sludge 147

RL29 Ravenna Sludge 2016 Sludge AK1 Akron 2016 Water AK2 Akron 2016 Water AK3 Akron 2016 Water RS6 Ravenna Source 2016 Water RS7 Ravenna Source 2016 Water RS8 Ravenna Source 2016 Water RS9 Ravenna Source 2016 Water RS10 Ravenna Source 2016 Water RS11 Ravenna Source 2016 Water AK12 7/7 Akron 2016 Water AK11 7/7 Akron 2016 Water AK10 7/7 Akron 2016 Water AK9 7/7 Akron 2016 Water AK8 7/7 Akron 2016 Water AK7 7/7 Akron 2016 Water AK6 7/7 Akron 2016 Water AK5 7/7 Akron 2016 Water AK4 7/7 Akron 2016 Water AK21 7/7 Akron 2016 Water AK20 7/7 Akron 2016 Water AK19 7/7 Akron 2016 Water AK18 7/7 Akron 2016 Water AK17 7/7 Akron 2016 Water AK16 7/7 Akron 2016 Water AK15 7/7 Akron 2016 Water AK14 7/7 Akron 2016 Water AK13 7/7 Akron 2016 Water AK22 7/7 Akron 2016 Water S-16-19 Sandusky 2016 Water S-16-20 Sandusky 2016 Water S-16-21 Sandusky 2016 Water S-16-22 Sandusky 2016 Water S-16-23 Sandusky 2016 Water S-16-24 Sandusky 2016 Water 148

S-16-25 Sandusky 2016 Water S-16-26 Sandusky 2016 Water S-16-27 Sandusky 2016 Water S-16-10 Sandusky 2016 Water S-16-11 Sandusky 2016 Water S-16-12 Sandusky 2016 Water S-16-13 Sandusky 2016 Water S-16-14 Sandusky 2016 Water S-16-15 Sandusky 2016 Water S-16-16 Sandusky 2016 Water S-16-17 Sandusky 2016 Water S-16-18 Sandusky 2016 Water S-16-1 Sandusky 2016 Water S-16-2 Sandusky 2016 Water S-16-3 Sandusky 2016 Water S-16-4 Sandusky 2016 Water S-16-6 Sandusky 2016 Water S-16-5 Sandusky 2016 Water S-16-7 Sandusky 2016 Water S-16-8 Sandusky 2016 Water S-16-9 Sandusky 2016 Water S-16-1 Sandusky 2016 Water S-16-2 Sandusky 2016 Water S-16-3 Sandusky 2016 Water S-16-4 Sandusky 2016 Water S-16-5 Sandusky 2016 Water S-16-6 Sandusky 2016 Water S-16-7 Sandusky 2016 Water S-16-8 Sandusky 2016 Water S-16-9 Sandusky 2016 Water S-16-10 Sandusky 2016 Water S-16-11 Sandusky 2016 Water S-16-12 Sandusky 2016 Water S-16-33 Sandusky 2016 Water S-16-34 Sandusky 2016 Water 149

S-16-35 Sandusky 2016 Water S-16-36 Sandusky 2016 Water S-16-37 Sandusky 2016 Water S-16-38 Sandusky 2016 Water S-16-39 Sandusky 2016 Water S-16-40 Sandusky 2016 Water S-16-41 Sandusky 2016 Water S-16-42 Sandusky 2016 Water S-16-43 Sandusky 2016 Water S-16-44 Sandusky 2016 Water S-16-45 Sandusky 2016 Water S-16-46 Sandusky 2016 Water S-16-47 Sandusky 2016 Water S-16-28 Sandusky 2016 Water S-16-29 Sandusky 2016 Water S-16-30 Sandusky 2016 Water S-16-31 Sandusky 2016 Water S-16-32 Sandusky 2016 Water

Table 8: Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Akron water treatment plant in the summer of 2018. Isolate Color Shape Elevation Opacity AK-W-23-18 White round Convex opaque AK-W-24-18 White round Convex opaque AK-W-25-18 White round Convex opaque AK-W-26-18 White round Convex opaque AK-W-27-18 White round Convex opaque AK-W-28-18 White round Convex opaque AK-W-29-18 White round Convex opaque AK-W-30-18 White round Convex opaque AK-W-31-18 White round Convex opaque 150

AK-W-32-18 White round Convex opaque AK-W-33-18 White round Convex opaque AK-W-34-18 White round Convex opaque AK-W-35-18 White round Convex opaque AK-W-36-18 White round Convex opaque AK-W-37-18 White round Convex opaque AK-W-38-18 White round Convex opaque AK-W-39-18 White round Convex opaque AK-W-40-18 White round Convex opaque AK-W-41-18 White round Convex opaque AK-W-42-18 White round Convex opaque AK-W-43-18 White round Convex opaque AK-W-44-18 White round Convex opaque AK-W-45-18 White round Convex opaque AK-W-46-18 White round Flat transparent AK-W-47-18 White round Flat transparent AK-W-48-18 White round Flat opaque AK-W-49-18 yellow / white round Convex opaque AK-W-50-18 White round Convex opaque AK-W-51-18 White round Convex opaque AK-W-52-18 White round Convex transparent AK-W-53-18 White round Convex opaque AK-W-54-18 yellow / white round Flat opaque AK-W-55-18 White round Flat opaque AK-W-56-18 White round Flat opaque AK-W-57-18 White round Flat opaque AK-W-58-18 White round Convex opaque

151

Table 9: Pure culture isolates and their colony characteristics as seen on R2A agar from the water and sludge samples collected from the Alliance water treatment plant in the summer of 2018.

Colony Color Shape Elevation Transparency AL-1-W-18 White irregular flat Opaque AL-2-W-18 White round/irregular flat Opaque AL-3-W-18 yellow/white round/irregular flat Opaque AL-4-W-18 yellow/white round/irregular flat Opaque AL-5-W-18 pink/white round flat Opaque AL-6-W-18 White round flat Opaque AL-7-W-18 White round flat Opaque AL-8-W-18 pink round flat Opaque AL-9-W-18 White round flat translucent AL-10-W-18 White round convex translucent AL-11-W-18 White round convex Opaque AL-12-W-18 White round convex Opaque AL-16-W-18 White pin-point flat translucent AL-17-W-18 yellow circle convex Opaque AL-18-W-18 White pin-point flat translucent AL-19-W-18 white/pink Circle flat Opaque AL-20-W-18 white/pink Circle convex Opaque AL-21-W-18 White pin-point flat translucent AL-22-W-18 White Circle convex translucent AL-23-W-18 yellow Circle convex Opaque AL-24-W-18 yellow/orange pin-point convex Opaque AL-25-W-18 yellow Circle convex Opaque AL-26-W-18 pink/white pin-point convex Opaque AL-27-W-18 White pin-point flat translucent AL-28-W-18 pink/white Circle convex translucent AL-29-W-18 pink pin-point convex translucent AL-30-W-18 white/cream Circle convex translucent AL-31-W-18 White circle w/ pin- flat translucent points AL-32-W-18 White pin-point flat translucent 152

AL-33-W-18 White pin-point convex Opaque AL-34-W-18 white/cream Circle convex Opaque AL-35-W-18 White pin-point flat translucent AL-36-W-18 yellow/orange pin-point flat translucent AL-37-W-18 White Circle flat translucent AL-38-W-18 white/cream Circle convex Opaque AL-39-W-18 light yellow pin-point flat translucent AL-40-W-18 orange pin-point flat Opaque AL-41-W-18 orange pin-point flat opaque AL-42-W-18 light yellow pin-point flat translucent AL-43-W-18 White Circle flat opaque AL-44-W-18 White Circle convex translucent AL-45-W-18 light yellow pin-point flat translucent AL-46-W-18 White Circle flat translucent AL-47-W-18 White Circle flat opaque

Note: tap water plates produced no colonies

AL-48-S-18 Clear no shape/veiny flat transparent AL-49-S-18 Cream pinpoint flat transparent AL-50-S-18 Cream pinpoint flat translucent AL-51-S-18 Cream pinpoint flat translucent AL-52-S-18 White no shape flat opaque AL-53-S-18 Cream Round flat translucent AL-54-S-18 Cream Round flat opaque AL-55-S-18 Cream Round flat translucent AL-56-S-18 Cream Round flat opaque AL-57-S-18 White irregular raised opaque AL-58-S-18 White Round flat opaque AL-59-S-18 Cream irregular flat translucent AL-60-S-18 Cream pinpoint flat translucent AL-61-S-18 Cream pinpoint flat translucent AL-62-S-18 Cream pinpoint flat translucent

153

AL-63-S-18 Cream Round flat translucent AL-64-S-18 White irregular flat opaque AL-65-S-18 Cream pinpoint flat translucent AL-66-S-18 Cream Round flat translucent AL-67-S-18 Cream pinpoint flat translucent AL-68-S-18 Cream pinpoint flat translucent AL-69-S-18 Cream pinpoint flat translucent AL-70-S-18 Cream pinpoint flat translucent

Table 10: Pure culture isolates and their colony characteristics as seen on R2A agar from the water and sludge samples collected from the Ravenna water treatment plant in the summer of 2018.

Colony Color Shape Elevation Transparency RA-1-S-18 white round flat Opaque RA-2-S-18 white round flat Opaque RA-3-S-18 white/cream round flat Opaque RA-4-S-18 white/cream stringy/round flat Translucent RA-5-S-18 white pin-point flat Opaque RA-6-S-18 white/cream round flat Translucent RA-7-S-18 white round raised Opaque RA-8-S-18 white/cream irregular/round flat Opaque RA-9-S-18 cream irregular/round flat Opaque RA-10-S-18 white/cream irregular/round flat Opaque RA-11-S-18 white/cream irregular flat Opaque RA-12-S-18 cream round raised Opaque RA-13-S-18 white/cream irregular flat Opaque RA-14-S-18 cream round raised Opaque RA-15-S-18 white pin-point convex Opaque RA-16-S-18 white pin-point/round convex Opaque RA-17-S-18 white pin-point/round convex Opaque RA-18-S-18 cream pin-point flat Translucent RA-19-S-18 white irregular/round raised Opaque RA-20-S-18 cream irregular/round raised Opaque RA-21-S-18 white irregular/round flat Opaque 154

RA-22-S-18 white pin-point/round flat Opaque RA-23-S-18 white irregular/round flat Opaque RA-24-S-28 white irregular/round raised Opaque RA-25-S-18 white stringy/round flat Opaque RA-26-S-18 white irregular flat Opaque RA-27-S-18 white irregular/round flat Opaque RA-28-S-18 cream round convex Opaque RA-29-S-18 white irregular/round flat Opaque RA-30-S-18 white irregular/round flat Opaque RA-31-S-18 yellow irregular flat Opaque

Note: "fished water" plates produced no colonies

RA-32-S-18 cream pinpoint flat Transparent RA-33-S-18 orange pinpoint flat Transparent RA-34-S-18 orange pinpoint flat Transparent RA-35-S-18 yellow pinpoint flat Transparent RA-36-S-18 orange pinpoint flat Transparent RA-37-S-18 orange pinpoint flat Transparent RA-38-S-18 white/cream pinpoint flat Transparent RA-39-S-18 cream pinpoint flat Transparent RA-40-S-18 cream pinpoint flat Transparent RA-41-S-18 cream/orange pinpoint flat Transparent RA-42-S-18 cream/orange round/pinpoint flat Transparent RA-43-S-18 cream round/pinpoint flat Transparent RA-44-S-18 cream round/pinpoint flat Transparent RA-45-S-18 white round/pinpoint flat Transparent RA-46-S-18 cream round/pinpoint flat Transparent RA-47-S-18 cream round/pinpoint flat Transparent RA-48-S-18 cream round flat Transparent RA-49-S-18 cream round/pinpoint flat Transparent RA-50-S-18 cream round/pinpoint flat Transparent RA-51-S-18 cream round/pinpoint flat Transparent

155

RA-52-S-18 orange round/pinpoint flat Transparent RA-53-S-18 cream round/pinpoint flat Transparent RA-54-S-18 orange round/pinpoint flat Transparent RA-55-S-18 cream round flat Transparent RA-56-W-18 cream round flat Transparent RA-57-W-18 clear no shape/veiny flat Transparent RA-58-W-18 white irregular/round convex Opaque RA-59-W-18 white irregular/round convex Opaque RA-60-W-18 white irregular/round convex Opaque RA-61-W-18 yellow pinpoint flat Opaque RA-62-W-18 cream round flat Translucent RA-63-W-18 cream round convex Opaque RA-64-W-18 cream round convex Translucent RA-65-W-18 cream round convex Opaque RA-66-W-18 orange irregular flat Transparent RA-67-W-18 orange irregular flat Transparent RA-68-W-18 cream irregular raised Opaque RA-69-W-18 orange irregular/round flat Transparent RA-70-W-18 orange irregular/round flat Transparent RA-71-W-18 orange irregular flat Transparent RA-72-W-18 white round raised Opaque RA-73-W-18 cream irregular flat Translucent RA-74-W-18 cream round flat Opaque RA-75-W-18 cream irregular flat Translucent RA-76-W-18 cream irregular flat Translucent RA-77-W-18 white round convex Opaque RA-78-W-18 cream pinpoint flat Transparent RA-79-W-18 yellow pinpoint flat Transparent RA-80-W-18 cream round flat Transparent RA-81-W-18 white round raised Opaque RA-82-W-18 white round raised Translucent RA-83-W-18 white round raised Opaque RA-84-W-18 cream irregular flat Translucent

156

RA-85-W-18 white irregular raised Opaque RA-86-W-18 cream round/pinpoint flat Transparent RA-87-W-18 cream round/pinpoint flat Transparent RA-88-W-18 white round raised Opaque RA-89-W-18 white irregular raised Opaque RA-90-W-18 cream round/pinpoint flat Transparent RA-91-W-18 white round/pinpoint flat Transparent RA-92-W-18 cream round/pinpoint flat Transparent RA-93-S-18 cream irregular flat Translucent RA-94-S-18 white pinpoint raised Opaque RA-95-S-18 white no shape/veiny flat Transparent RA-96-S-18 cream pinpoint raised Opaque RA-97-S-18 white irregular flat Opaque RA-98-S-18 white irregular flat Opaque RA-99-S-18 white irregular flat Opaque RA-100-S-18 cream round/pinpoint flat Translucent RA-101-S-18 white irregular flat Opaque RA-102-S-18 cream round flat Translucent RA-103-S-18 white irregular flat Opaque RA-104-S-18 cream round flat Translucent RA-105-S-18 cream round flat Translucent RA-106-S-18 white irregular flat Opaque RA-107-S-18 cream round flat Translucent RA-108-S-18 cream round flat Translucent RA-109-W-18 white irregular raised Opaque RA-110-W-18 white round raised Opaque RA-111-W-18 cream irregular flat Opaque RA-112-W-18 white pinpoint raised Opaque RA-113-W-18 white pinpoint flat Translucent RA-114-W-18 white irregular raised Opaque RA-115-W-18 white pinpoint flat Opaque RA-116-W-18 white irregular raised Opaque RA-117-W-18 cream round/pinpoint flat Transparent

157

RA-118-W-18 cream pinpoint flat Transparent RA-119-W-18 yellow round/pinpoint flat Opaque RA-120-W-18 cream round flat Translucent RA-121-W-18 cream round raised Opaque RA-122-W-18 cream round flat Opaque RA-123-W-18 cream pinpoint flat Translucent RA-124-W-18 yellow pinpoint flat Opaque RA-125-W-18 cream round/pinpoint flat Translucent RA-126-W-18 cream round/pinpoint flat Transparent RA-127-W-18 white pinpoint flat Opaque RA-128-W-18 cream round/pinpoint raised Opaque RA-129-W-18 cream round flat Translucent RA-130-W-18 cream/clear no shape flat Transparent RA-131-W-18 cream irregular flat Transparent RA-132-W-18 cream round raised Opaque RA-133-W-18 cream pinpoint flat Translucent RA-134-W-18 cream pinpoint flat Translucent RA-135-W-18 cream pinpoint flat Translucent RA-136-W-18 cream pinpoint flat Translucent RA-137-S-18 cream pinpoint flat Translucent RA-138-S-18 cream round/pinpoint flat Opaque RA-139-S-18 cream round flat Translucent RA-140-S-18 cream round flat Translucent RA-141-S-18 cream round raised Translucent RA-142-S-18 white irregular flat Opaque RA-143-S-18 cream round raised Opaque RA-144-S-18 cream round flat Translucent RA-145-S-18 cream round flat Translucent RA-146-S-18 white irregular flat Opaque RA-147-S-18 cream round/pinpoint flat Translucent RA-148-S-18 cream round flat Translucent RA-149-S-18 orange pinpoint convex Transparent RA-150-S-18 cream round convex Translucent

158

RA-151-S-18 orange pinpoint convex Transparent RA-152-S-18 cream round convex Translucent RA-153-S-18 white pinpoint flat Transparent RA-154-S-18 white pinpoint flat Translucent RA-155-S-18 white round/pinpoint flat Translucent RA-156-S-18 white round/pinpoint flat Translucent

Table 11: Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Kent water treatment plant in the summer of 2018.

Colony Color Shape Elevation Transparency KW-1-W-18 White round convex Opaque KW-2-W-18 yellow round flat Translucent KW-3-W-18 White round convex Translucent KW-4-W-18 White round flat Translucent KW-5-W-18 yellow round convex Translucent KW-6-W-18 White round convex Opaque KW-7-W-18 yellow/white round convex Opaque KW-8-W-18 White round convex Translucent KW-9-W-18 yellow/white round convex Translucent KW-10-W-18 yellow/white round convex Opaque KW-11-W-18 White round flat Opaque KW-12-W-18 yellow/white round convex Opaque KW-13-W-18 yellow/white round convex Opaque KW-14-W-18 yellow round convex Opaque KW-15-W-18 Pink round flat Translucent KW-16-W-18 yellow round flat Translucent KW-17-W-18 orange round convex Translucent KW-18-W-18 cream pin-point/round flat Translucent KW-19-W-18 yellow pin-point/round convex Opaque KW-20-W-18 yellow pin-point/round convex Opaque KW-21-W-18 cream pin-point/round flat Translucent 159

KW-22-W-18 White pin-point flat Translucent KW-23-W-18 White pin-point flat Translucent KW-24-W-18 White pin-point flat Translucent KW-25-W-18 White pin-point flat Translucent KW-26-W-18 White pin-point flat Translucent KW-27-W-18 White pin-point flat Translucent KW-28-W-18 White pin-point flat Translucent KW-29-W-18 White pin-point flat Translucent KW-30-W-18 cream irregular/round flat Opaque KW-31-W-18 cream irregular/round flat Opaque KW-32-W-18 White irregular/round convex Opaque KW-33-W-18 White pin-point/round flat Opaque KW-34-W-18 cream irregular flat Translucent KW-35-W-18 yellow round flat Translucent KW-36-W-18 White pin-point flat Translucent KW-37-W-18 White pin-point flat Translucent KW-38-W-18 White pin-point flat Translucent KW-39-W-18 yellow pin-point/round raised Translucent KW-40-W-18 White pin-point/round convex Opaque KW-41-W-18 cream round flat Translucent KW-42-W-18 cream pin-point flat Translucent KW-43-W-18 White pin-point flat Translucent KW-44-W-18 White pin-point flat Translucent KW-45-W-18 cream pin-point flat Translucent KW-46-W-18 White pin-point flat Translucent KW-47-W-18 cream pin-point flat Translucent KW-48-W-18 cream pin-point flat Translucent KW-49-W-18 cream irregular flat Translucent KW-50-W-18 White pin-point convex Opaque KW-51-W-18 cream irregular flat Translucent KW-52-W-18 cream pin-point flat Translucent KW-53-W-18 cream pin-point flat Translucent KW-54-W-18 cream pin-point flat Translucent

160

KW-55-W-18 cream pin-point flat Translucent KW-56-W-18 cream pin-point flat Translucent KW-57-W-18 cream irregular flat Translucent KW-58-W-18 cream pin-point convex Translucent KW-59-W-18 cream pin-point flat Translucent KW-60-W-18 cream pin-point flat Translucent KW-61-W-18 cream pin-point flat Translucent KW-62-W-18 cream pin-point flat Translucent KW-63-W-18 cream pin-point convex Opaque KW-64-W-18 cream pin-point flat Translucent KW-65-W-18 cream round flat Transparent KW-66-W-18 cream irregular flat Translucent KW-67-W-18 White round flat Translucent KW-68-W-18 yellow/cream round flat Translucent KW-69-W-18 cream round flat Transparent KW-70-W-18 brown round flat Transparent KW-71-W-18 cream round flat Transparent KW-72-W-18 White irregular/round convex Translucent

Table 12: Pure culture isolates and their colony characteristics as seen on R2A agar from the water samples collected from the Sandusky water treatment plant in the summer of 2018. Colony Color Shape Elevation Transparency SA-1-W-18 cream/yellow pinpoint flat Opaque SA-2-W-18 cream pinpoint flat Translucent SA-3-W-18 cream/yellow pinpoint flat Opaque SA-4-W-18 cream/yellow pinpoint flat Opaque SA-5-W-18 cream/yellow circle flat Opaque SA-6-W-18 cream/yellow pinpoint flat Opaque SA-7-W-18 cream/yellow circle flat Opaque SA-8-W-18 cream/yellow pinpoint flat Opaque SA-9-W-18 cream pinpoint flat Opaque SA-10-W-18 cream pinpoint flat Opaque 161

SA-11-W-18 SA-12-W-18 yellow irregular/round flat Opaque SA-13-W-18 cream circle flat Translucent SA-14-W-18 cream pinpoint flat Opaque SA-15-W-18 cream irregular/circle flat Opaque SA-16-W-18 yellow circle flat Opaque SA-17-W-18 cream circle convex Opaque SA-18-W-18 cream pinpoint flat Opaque SA-19-W-18 cream pinpoint/circle flat Opaque SA-20-W-18 SA-21-W-18 cream circle convex Opaque SA-22-W-18 cream circle convex Opaque SA-23-W-18 cream circle flat Opaque SA-24-W-18 cream circle flat Opaque SA-25-W-18 cream pinpoint flat Opaque SA-26-W-18 cream pinpoint flat Translucent SA-27-W-18 cream circle flat Opaque SA-28-W-18 cream pinpoint/circle flat Opaque SA-29-W-18 yellow irregular/circle flat Opaque SA-30-W-18 cream circle convex Translucent SA-31-W-18 white/cream pinpoint flat Translucent SA-32-W-18 yellow irregular/circle flat Opaque SA-33-W-18 cream pinpoint flat Translucent SA-34-W-18 cream/yellow irregular raised Opaque SA-35-W-18 yellow irregular/circle flat Translucent SA-36-W-18 light yellow irregular/circle flat Translucent SA-37-W-18 white/cream pinpoint/irregular flat Translucent SA-38-W-18 cream/yellow circle flat Translucent SA-39-W-18 yellow circle w/ edges flat Translucent SA-40-W-18 cream circle / pinpoint flat Translucent SA-41-W-18 yellow/white circle w/ edges flat Translucent SA-42-W-18 cream pinpoint flat Translucent SA-43-W-18 yellow/white pinpoint w/ edges flat Translucent

162

SA-44-W-18 yellow/cream circle w/ edges flat Translucent SA-45-W-18 white/cream circle w/ edges flat Translucent SA-46-W-18 cream pinpoint/circle flat Translucent SA-47-W-18 cream circle flat Translucent SA-48-W-18 cream circle flat Opaque SA-49-W-18 yellow circle convex Opaque SA-50-W-18 light yellow circle convex Opaque SA-51-W-18 light yellow circle convex Opaque SA-52-W-18 light yellow circle convex Opaque SA-53-W-18 yellow pinpoint flat Translucent SA-54-W-18 yellow/cream circle flat Translucent SA-55-W-18 cream circle convex Translucent SA-56-W-18 cream irregular flat Translucent SA-57-W-18 light yellow circle convex Opaque SA-58-W-18 light yellow circle convex Opaque SA-59-W-18 light yellow circle convex Opaque SA-60-W-18 light yellow circle convex Opaque SA-61-W-18 cream circle convex Opaque SA-62-W-18 bright yellow irregular/circle flat Opaque SA-63-W-18 cream circle convex Opaque SA-64-W-18 cream irregular/circle flat Opaque SA-65-S-18 cream round w/ rough edges flat Opaque SA-66-S-18 cream irregular convex Opaque SA-67-S-18 cream round raised Opaque SA-68-S-18 cream round/stringy/spiral flat Opaque SA-69-S-18 white round raised Transparent SA-70-S-18 white stringy/spiral flat Opaque SA-71-S-18 white round convex Translucent SA-72-S-18 cream irregular raised Opaque SA-73-S-18 cream irregular w/ rough edges flat Translucent SA-74-S-18 white irregular w/ rough edges flat Translucent SA-75-S-18 cream irregular w/ rough edges flat Translucent SA-76-S-18 cream round/stringy/spiral flat Opaque

163

SA-77-S-18 white round raised Opaque SA-78-S-18 white/cream round/stringy/spiral convex Opaque SA-79-S-18 cream round/stringy/spiral convex Opaque SA-80-S-18 white round w/ rough edges flat Opaque SA-81-S-18 white round/stringy/spiral flat Opaque SA-82-S-18 cream round convex Opaque SA-83-S-18 white round convex Transparent SA-84-S-18 clear round convex Opaque SA-85-S-18 cream round/stringy/spiral convex Opaque SA-86-S-18 white round convex Opaque SA-87-S-18 white round/stringy/spiral flat Opaque SA-88-S-18 white & pink round/stringy/spiral convex Opaque SA-89-S-18 white irregular w/ rough edges flat Opaque SA-90-S-18 white irregular w/ rough edges flat Opaque SA-91-S-18 white/cream pinpoint/irregular flat Translucent SA-92-S-18 white round w/ rough edges flat Opaque SA-93-S-18 white pinpoint flat Transparent SA-94-S-18 white round w/ rough edges flat Translucent SA-95-S-18 white round w/ rough edges flat Opaque SA-96-S-18 white round flat Opaque SA-97-S-18 white round/stringy flat Translucent SA-98-S-18 yellow pinpoint convex Opaque SA-99-S-18 white round w/ rough edges flat Translucent SA-100-S-18 cream/orange round convex Opaque SA-101-S-18 cream round convex Opaque SA-102-S-18 cream round convex Opaque SA-103-S-18 cream round convex Opaque SA-104-S-18 cream irregular w/ rough edges flat Translucent SA-105-S-18 white irregular flat Opaque SA-106-S-18 cream/orange round convex Opaque SA-107-S-18 white pinpoint flat Translucent SA-108-S-18 yellow round convex Opaque

164