Sustainable Methods for Cyanotoxin Treatment and Discovery of the Cyanophage THESIS Presented in Partial Fulfillment of the Requ

Sustainable Methods for Cyanotoxin Treatment and Discovery of the Cyanophage

THESIS

Presented in Partial Fulfillment of the Requirements for the Degree Master of Science in the Graduate School of The Ohio State University

Xuewen Jiang

Graduate Program in Food Science and Technology

The Ohio State University

2017

Master's Examination Committee:

Jiyoung Lee, Advisor

Ahmed Yousef

Luis Rodriguez-sonata

Copyrighted by

Xuewen Jiang

2017

Abstract

Cyanobacterial harmful algal bloom (CHABs) has been a great concern due to the detrimental effects on ecosystem, animal and human health. Overgrowth of CHABs causes hypoxia and some species produce unpleasant odor compounds. More importantly, multiple toxins (cyanotoxins) produced by CHABs can be harmful to animal and human health by damaging internal organs, including liver, kidney, reproductive and neurological systems. Water-mediated activities are believed to be the common exposure routes for human to these toxins. Many studies reported that these toxins can be accumulated into food or dietary supplements via contaminated water, since the guideline for agriculture and aquaculture water use is not established yet. Therefore, strategies for controlling both CHABs and their toxins are important and imperative for protecting environmental and food safety. Chapter 1 summarized the literature review about CHABs and their toxins, and identified knowledge gaps. Furthermore, previous research regarding to cyanophages and their ecological roles are also reviewed because understanding of cyanophage, natural predators of cyanobacteria, is critical to understanding the bloom formation and evolution.

Among the cyanotoxins, the most abundant cyanotoxin is microcystin (MC), which is a seven-amino-acid peptide, consisting of more than 150 congeners. MC shows toxicity via inhibiting protein phosphatase 1 and 2A, and generate reactive oxygen species (ROS), leading to cell apoptosis in the involved organs. MCs are chemically stable against ii sunlight, extreme pH and boiling, therefore, it is difficult to treat the contaminated water once MCs are released. Physical, chemical and biological methods have been applied to treat MCs in water, but few of them can be successfully applied for MC- contaminated food. New technologies for MC treatment are proposed in Chapter 2. The objective of

Chapter 2 was to develop sustainable and non-chemical-based methods for controlling

MCs: 1) ultraviolet with TiO2, which is a common food additive as well as a photocatalyst; and 2) cold plasma, which is a non-thermal treatment using ionized gas.

Natural MCs were extracted from Microcystis aeruginosa and treated with several combinations: 1) UV at intensity of 1470 μW/cm2 [high] or 180 μW/cm2 [low]; 2) cold plasma; and 3) no treatment in a dark room (control). To determine synergistic effects, nanoparticles (TiO2) were coated on the outside of the UV treatment chamber prior to irradiation. The MC degradation efficiency was enhanced by the reusable TiO2 coating at lower UV intensity by 10 percent, but no significant difference was observed at higher intensity of UV. Cold plasma removed MCs rapidly (80% and 92% in 1 and 2 hours, respectively) under experimental conditions, indicating that it can be easily and practically used in household and industrial setting.

In environmental settings, controlling CHABs formation is a better and effective way to reduce toxin production. Many previous studies have focused on environmental factors on blooms, but the role of cyanophage (viruses whose hosts are cyanobacteria) in the ecology of toxin production is much less understood, especially in freshwater.

Cyanophage are known to contribute to biological controlling of CHABs by influencing the metabolism and evolution of cyanobacteria. Therefore, the information on the

iii characteristics of cyanophage infection is crucial in potential bloom control. In Chapter 3, findings about cyanophage isolated from Lake Erie during bloom season were reported.

The objectives of this study are to 1) isolate and characterize cyanophages from Lake

Erie; and 2) examine the host-cyanophage interactions using multiple tools, especially atomic force microscopy (AFM). Cyanophage isolates were inoculated into toxin- producing Microcystis aeruginosa (host) originated from Lake Erie. The dynamic interactions between phages and hosts were monitored using spectrophotometer, fluorimeter, quantitative PCR and AFM. The structural and genetic characters of the cyanophages were identified by using transmission electron microscope (TEM) and PCR.

The Podoviridae ~300 nm in size were infectious against the toxic strain of M. aeruginosa. The growth and photosynthesis capacity of hosts was significantly inhibited after cyanophage infection and cellular damages on membranes over time were clearly observed. We also found that UV irradiation with proper dose can induce the cyanophage shifting from their lysogenic stage to lytic stage. The psbA (photosynthesis core protein

A) and the gp58 (special capsid protein) genes were identified, which showed high similarities of the same genes identified in marine cyanophages, indicating a close evolutionary root.

In summary, this thesis reports comprehensive knowledge about CHABs and their toxins. In addition, emerging techniques were applied in this study to provide informative and essential basis to develop controlling strategies in various settings when toxins are present.

Acknowledgments

I’d like to firstly express my great appreciation to my advisor, Dr. Jiyoung Lee.

She provides continuous guidance from the beginning of my admission no matter in my academic or personal life. Without her trusts and supports, I cannot finish my research and thesis.

I appreciate all the help from my lab member Seungjun Lee, who answered my questions patiently, taught me when I made mistakes and comforted me when I was upset. It is great to work with him during two years. I also feel grateful to all my lab members and my committee members. They always provide essential help without hesitates and make me feel at home.

Thanks for all the professors and friends. Your supports are the best gift.

Last but not least, I appreciate all the supports from my family. They are my motivation to move forward, as well as my solid backup. Thank you for letting me choose my own life and supporting me all the time.

Vita

September 2008 to June 2011 ...... Shanxi Experimental Secondary School

August 2011 to July 2015 ...... B.S. Biological Science, Nanjing University

August 2015 to present ...... M.S. Department of Food Science and

Technology, The Ohio State University

Publications

Lee J, Lee S, Jiang X. Cyanobacterial Toxins in Freshwater and Food: Important Sources of Exposure to Humans. Annual Review of Food Science and Technology. 2017

Apr;8(1).

Fields of Study

Major Field: Food Science and Technology

Table of Contents

Abstract ...... ii

Acknowledgments...... v

Vita ...... vi

Publications ...... vi

Fields of Study ...... vi

Table of Contents ...... vii

List of Tables ...... xi

List of Figures ...... xiii

Chapter 1: literature review ...... 1

1. Cyanotoxins: a growing concern for food safety and public health ...... 1

1.1 Toxin-producing cyanobacteria: Harmful algal blooms ...... 1

1.1.1 Introduction of cyanobacteria ...... 1

1.1.2 Concerns about cyanobacterial blooms ...... 2

1.1.3 Factors affecting bloom formation ...... 4

1.1.4 Monitoring of blooms ...... 6

vii

1.2 Microcystins: the most widespread cyanotoxin ...... 7

1.2.1 Introduction to microcystins ...... 7

1.2.2 Health impacts and guidelines of microcystins ...... 8

1.2.3 Exposure routes of microcystins for humans ...... 13

1.2.4 Effects of MCs on agriculture ...... 19

1.2.5 Quantification of microcystins ...... 21

1.2.6 Decontamination of MCs...... 22

1.3 Health concerns from other cyanotoxins ...... 27

2. Cyanophages-cyanobacteria interplay ...... 29

2.1 Bacteriophage-bacteria interaction ...... 29

2.1.1 Introduction of virus with the emphasis of bacteriophage ...... 29

2.1.2 Co-evolution of bacteriophages and their hosts: an arms-race ...... 32

2.1.3 Biocontrol of foodborne pathogens using bacteriophages ...... 33

2.2 Cyanophages ...... 36

Chapter 2: Sustainable Methods for Decontamination of Microcystin in Water using Cold

Plasma and UV with Reusable TiO2 Nanoparticle Coating ...... 41

1. Abstract ...... 41

2. Introduction ...... 42

3. Materials and Methods ...... 45

viii

3.1 MC preparation ...... 45

3.2 MC treatments and measurement ...... 45

3.3 Statistical Analysis ...... 46

4. Results ...... 47

4.1. Synergistic Effects of TiO2 with UV on MC degradation ...... 47

4.2. Effectiveness of Cold plasma on MC degradation ...... 47

4.3 Kinetic analysis...... 48

5. Discussion ...... 49

6. Conclusions ...... 52

7. Figures ...... 54

8. Tables ...... 57

Chapter 3: Discovery of lytic cyanophages and their interaction mechanisms with toxic cyanobacteria: A potential biological control of harmful algal blooms ...... 59

1. Abstract ...... 59

2. Introduction ...... 60

3. Material and methods ...... 62

3.1 Collection, concentration and screening of lytic cyanophage ...... 62

3.2 TEM ...... 63

3.3 Presence of special genes using PCR ...... 63

3.4 The effects of cyanophage infections on Microcystis aeruginosa...... 64

3.5 AFM...... 65

3.6 UV-induced activation of the prophage...... 65

3.7 Statistical analysis...... 66

4. Results ...... 66

4.1 Screening for lytic cyanophage ...... 66

4.2 TEM ...... 66

4.3 Presence of psbA and gp58 genes ...... 66

4.4 The effects of Ma-LEP infection on M. aeruginosa ...... 67

4.5 AFM...... 68

4.6 UV-induced activation of the prophage...... 68

5. Discussion ...... 69

6. Figures ...... 74

7. Tables ...... 81

8. Supplementary Data ...... 84

References ...... 86

List of Tables

Table 1.1 Representative freshwater cyanobacteria...... 2

Table 1.2 Guidelines of microcystins in drinking water in different countries...... 11

Table 1.3 Guidelines for recreational water (agricultural water)...... 12

Table 1.4 Bioaccumulation and physiological changes of crops and seafood exposed to

MCs ...... 15

Table 1.5 Toxicity and health effects of other cyanotoxins...... 28

Table 1.6 Bacterial virulence properties altered by bacteriophages...... 33

Table 1.7 Examples of freshwater cyanophages...... 37

Table 2.1 Degradation rate constants (k) and half-lives (t½) in the first- or second-order reaction kinetics in microcystin removal by UV and UV with TiO2...... 57

Table 2.2 Degradation rate constants (k) and half-lives (t½) in the first- or second-order reaction kinetics in microcystin removal by cold plasma...... 58

Table 2.3 Energy per order (EEO) of each treatment...... 58

Table 3.1 PCR condition and primer sequences used in this study ...... 81

Table 3.2 The growth/ pigment production of M. aeruginosa with or without Cyanophage

LEP infection as a function of time in a linear model...... 83

Table 3.3 The growth/ pigment production of M. aeruginosa with or without lysogenic

Cyanophage LEP infection after UV irradiation as a function of time in a linear model. 83 xi

Table 3.4 Sampling sites in Lake Erie...... 84

Table 3.5 Gene sequences of host and Cyanophage LEP...... 85

xii

List of Figures

Figure 1.1 Toxin exposure scenarios near freshwater environments thataffected by cyanobacterial blooms...... 4

Figure 1.2 Summary of known feedback mechanisms between climate change and bloom formation...... 5

Figure 1.3 The structure of microcystins. X, Z, R1 and R2 varies in different congeners. . 8

Figure 1.4 The effects of MC-LR and CYN on plant growth, development and biochemical processes and human health risks ...... 20

Figure 1.5 Major principle of ADDA-based ELISA ...... 22

Figure 1.6 Schematic diagram of photocatalytic mechanism of TiO2...... 24

Figure 1.7 Pathway of MC biodegradation...... 26

Figure 1.8 Diagram of bacteriophage...... 30

Figure 1.9 Life cycle of bacteriophages...... 31

Figure 1.10 WHO estimated global foodborne disease burden...... 34

Figure 1.11 Model of the metabolic pathways regulated by cyanophage host-like genes in infected cyanobacterial cells...... 38

Figure 2.1 Schematic diagram of MC treatment in this study...... 54

Figure 2.2 The MC degradation rate in water treated with UV and UV/TiO2...... 55 xiii

Figure 2.3 The MC degradation rate in water treated with cold plasma...... 56

Figure 3.1 TEM images of repeating cyanophages with short tail and special capsids. ... 74

Figure 3.2 Phylogenic tree of Cyanophage LEP based on the sequence of psbA gene .... 75

Figure 3.3 The dynamic changes of M. aeruginosa infected by Cyanophage LEP in two weeks...... 76

Figure 3.4 Scenario of cell lysis of M. aeruginosa caused by Cyanophage LEP infection. .

...... 77

Figure 3.5 Induction of lysogenic Cyanophage LEP by UV irradiation...... 79

Figure 3.6 Color change of host culture...... 84

Figure 3.7 Bands of corresponding genes on 2% argarose gel...... 84

xiv

Chapter 1: literature review

1. Cyanotoxins: a growing concern for food safety and public health

1.1 Toxin-producing cyanobacteria: Harmful algal blooms

1.1.1 Introduction of cyanobacteria

Toxin-producing, harmful, cyanobacteria algal blooms mainly refer to the overgrowth of toxin-producing cyanobacteria (commonly known as blue-green algae), including Microcystis spp., Anabaena spp., Nodularia spp. and others (Table 1.1), which usually occur in warm, static water (Cheung et al., 2013; Paerl et al., 2001). With similar characteristics of algal cells, such as photosynthesis, green color, and relatively large size

(0.4-40 μm), cyanobacteria have been considered as algae for a long time (Whitton &

Potts, 2007). However, new genetic evidence and morphological findings have revealed that their phylogenic nature is prokaryotic (Alexandrov, 2016). Cyanobacteria are the oldest organisms in the world (~3.5 billion years) without defined nuclei and organelles, and most of them are Gram-negative (Hoiczyk & Hansel, 2000; Knoll, 2015). In the history of evolution, cyanobacteria play a crucial role in producing atmospheric oxygen

(anaerobic photosynthesis) and in the nitrogen circle (atmosphere N2 fixation by nitrogenase) (Shaw, 2016). Moreover, they can survive under phosphorus (P)-limited

1 conditions by using P stored during P-enriched seasons, which allows them to adapt to most ecosystems (Wiegand & Pflugmacher, 2005).

Table 1.1 Representative freshwater cyanobacteria (modified from Paerl et al. 2001 and Watson et al. 2016). Genus Metabolites Preferred Bloom Conditions Anabaena Anatoxin-a, Anatoxin-a(s), P-enriched, warm, microcystins, saxitoxins, stratified, long- BMAA; Geosmin, MIB. residence time, high Aphanizomenon Anatoxin-a, Anatoxin-a(s), irradiance, eutrophic. Cylindrospermopsin, microcystins, saxitoxins, LPS; Geosmin. Cylindrospermopsis Cylindrospermopsin, saxitoxins, BMAA. Nodularia Nodularins. Microcystis Microcystins, BMAA. N- and P-enriched, Planktothrix &Oscillatoria Anatoxin-a, Anatoxin-a(s), eutrophic conditions, microcystins, Geosmin, warm, stratified, long LPS, BMAA; MIB. residence time.

MIB: 2-methylisoborneol; BMAA: β-Methylamino-L-alanine; LPS: lipopolysaccharide.

1.1.2 Concerns about cyanobacterial blooms

Toxin-producing and non-toxin-producing strains of cyanobacteria coexist in the environment, and the factors triggering toxin productions are unclear (Cheung et al.,

2013; Yu et al., 2014). Still, cyanobacterial blooms have been a growing concern in the

United States and globally, with increasing frequency, duration, and intensity (Davis &

Gobler, 2016). Figure 1.1 summarizes the potential effects of CHABs on human/animal activities and related environments.

First, blooms can produce problematic toxins including microcystins (MCs), nodularins, cylindrospermopsin and others (Table 1.1). These toxins pose threats to human health and to other living organisms. For example, MCs are hepatotoxins (causing liver damage) and anatoxin-a is a neurotoxin (Van Apeldoorn et al., 2007). Cyanotoxins can enter other environments like soil, groundwater, and the food web. Therefore, toxins are not limited to bloom-affected water bodies, but are widely distributed throughout the earth (Funari & Testai, 2008; Lee et al., 2017; Machado et al., 2016a). In addition, the overgrowth of blooms can damage or clog the gills of fish or deplete the oxygen in a water body, leading to a reduction in biodiversity or even creating “dead zones” during bloom seasons (Zingone & Oksfeldt Enevoldsen, 2000). Some species may also produce compounds affecting taste-and-odor, such as geosmin and 2-methylisoborneol (MIB), that affect local people and tourists (Zamir et al. 2001). Each year, the cyanobacterial blooms cause millions of dollars in loss to tourism, fishing industries, and to management (Hoagland et al. 2002).

Figure 1.1 Toxin exposure scenarios near freshwater environments thataffected by cyanobacterial blooms. (From Lee et al. 2017)

1.1.3 Factors affecting bloom formation

There are many conditions that favor bloom formation. The eutrophication of water bodies is believed to be a major factor: scientists believe that P influx, both from non- point sources (e.g., agricultural runoff) and point sources (e.g., wastewater discharge, municipal or industrial loading), accelerates the growth of cyanobacteria and promote the growth of the toxic over the non-toxic strain (Anderson et al., 2002; Bertani et al., 2016;

Paerl et al., 2015). The ideal ratio of N to P is less than 15 to 1 (Paerl et al., 2001).

Another contributor is climate change; the composition of blooms may be affected due to changes in environmental or biological factors (Glibert et al., 2014). Anderson et al. (2012) have summarized the comprehensive factors that have been involved in bloom formation (Figure 1.2). Besides the increased water temperature, elevated atmospheric

CO2 promotes photosynthesis and changes the pH of water bodies to conditions that are

4 favorable for bloom formation. The food web structure, such as the abundance of grazers, may shift in response to climate change, leading to less predation of cyanobacteria. In addition, a higher frequency of storms may bring more nutrients into the receiving water, leading to more eutrophication and higher bloom frequency.

Figure 1.2 Summary of known feedback mechanisms between climate change and bloom formation. (modified from Anderson et al. 2012).

1.1.4 Monitoring of blooms

To monitor blooms, the World Health Organization (WHO) established guidelines for recreational water using chlorophyll-a, cyanobacterial cell counts, and microcystins as indicators of health risk (WHO, 2003b), but these indicators reflect inconsistent risk levels (EPA, 2016c). However, the measurement of cyanotoxins or cell density using molecular tools (e.g., qPCR) takes a relatively long time, therefore more rapid methods are needed for monitoring. Phycocyanin, another essential pigment produced by cyanobacteria, has been recommended by the United States Environmental Protection

Agency (EPA 2016b) as an indicator of the presence of cyanobacterial blooms. In vivo phycocyanin measurements (observing the peak absorbance at 620 nm) have been developed, which show good correlation with cell densities and microcystin concentrations (Hunter et al., 2016; Makarewicz et al., 2009; Marion et al., 2012;

Salmaso et al., 2016). Furthermore, this approach enables remote sensing, by using satellite images with a MERIS (Medium Resolution Imaging Spectrometer) to monitor dynamic changes over time of blooms in higher scales (Lee et al., 2015). USGS (2017) reported that “two USGS-operated Landsat satellites and a new European Space Agency satellite” have been recruited to collect satellite data. However, one limitation of this method is that images from satellites can be easily affected by poor atmospheric conditions. Thus, the USEPA, NASA, NOAA, and USGS initiated an “early warning indicator system” in 2015, called the Cyanobacteria Assessment Network (CyAN). One objective is to adopt “second derivative spectral shape algorithms” to overcome this disadvantage. Another disadvantage is that the minimum resolution is 300×300 meters

6 per pixel (EPA et al., 2015), which misses most small lakes and ponds. Dr. Jiyoung Lee has been working on multi-spectral scanning the color profile using unmanned aerial vehicles over lakes that are used for drinking and irrigation water source. This approach may be feasible to provide an early warning in smaller areas (Heidersbach et al., 2016).

1.2 Microcystins: the most widespread cyanotoxin

1.2.1 Introduction to microcystins

Cyanotoxins are secondary metabolites of cyanobacteria. Microcystins (MCs) are the most abundant and widely distributed cyanotoxin in freshwater produced by multiple cyanobacteria (Table 1.1) (Carrasco et al., 2006; Su et al., 2015). MCs are cyclic seven- amino-acid peptides, including the Mdha (N-methyldehydroalanine) at position 7 and the

ADDA [(2S,3S,4E,6E,8S,9S)-3-amino-9-methoxy-2,6,8-trimethyl-10-phenyldeca-4,6- dienoic acid] at position 5, which is a unique structure contributing to their toxicity

(Carmichael et al., 1988; Catherine et al., 2013; Dawson, 1998). MCs contain more than

150 congeners due to a variation in position 2 (X), 3(R1), 4(Y), and 7(R2) (Figure 1.3), and each variant has shown different properties (Foss & Aubel, 2015; Valério et al.,

2016). For example, MC-LR (X=Leu, Y=Arg, R1=R2=CH3) is the most well-known and thoroughly studied MC with the highest toxicity (Gehringer et al., 2005), while MC-RR

(X= Y= Arg, R1 =R2 = CH3) is the most hydrophilic (EPA, 2015b; Tsuji et al., 2001).

Figure 1.3 The structure of microcystins. X, Z, R1 and R2 varies in different congeners.

1.2.2 Health impacts and guidelines of microcystins

MCs can cause acute and chronic toxicity in mammals, birds, amphibians, and fish, including gastrointestinal inflammation, pneumonia, dermatitis, and multi-organ damage (e.g., liver, kidney, neural system, reproductive system, etc.), even liver failure or death (Boopathi & Ki, 2014; Calomeni & Rodgers, 2015; Cheung et al., 2013). Long- term studies show that MCs are a potential tumor-promoter. The lethal dose for 50% mortality (LD50) of MCs ranges from 45 to 1000 μg/ kg body weight (WHO, 2003b). As a reference, the highest toxicity of aflatoxin B1 (AFB1) is 300 μg/ kg body weight

(Newberne & Butler, 1969).

MCs can be actively transported via the bile acid carrier transport system, which is widely expressed, leading to severe liver damage. Transportation can be inhibited by other competitive substances (e.g., bile salts cholate, taurocholate), or non-competitively

8 by antamanide, rifampicin, etc. (Feurstein et al., 2010; Wiegand & Pflugmacher, 2005).

MCs inactivate serine/threonine protein phosphatases (type 1 and 2A, PP-1 and PP2A) by introducing an ADDA group into their catalytic sites (Campos & Vasconcelos, 2010;

Valério et al., 2016). Structural changes in ADDA lead to less toxicity of MCs against

PP1 and PP2A (Wiegand & Pflugmacher, 2005). PP1 and PP2A are associated with the balance of cellular phosphorylation, which is important for the dynamics of microtubules, neurofilaments and actin cytoskeletons, therefore the inactivation of PP1 and PP2A leads to cytoskeletal disorganization and cellular disruption (Hoffman et al. 2017). In addition,

MCs induce oxidative stress by generating reactive oxygen species (ROS) that cause

DNA damage and cellular lipid oxidation, which further leads to the loss of membrane integrity (Chen et al., 2016; Valério et al., 2016). Finally, cell apoptosis is inevitable in the involved organs. It has also been reported that MCs affect PP2A expression and damage the hypothalamic-pituitary-gonad (HPG) axis with sex-dependent effects (Chen et al., 2016; Liu et al., 2016c; Valério et al., 2016).

Different countries have applied various guidelines for MCs (Table 1.2). The WHO

(2003a) considers MCs below 1 part per billion (ppb) safe for consumption in drinking water, which is widely accepted by most countries. The USEPA (2015a) recently published new guidelines stating that MCs below 0.3 ppb in drinking water are safe for pre-school age children, and levels below 1.6 ppb are safe for other populations.

However, while agricultural water use generally follows recreational water guidelines, it is not clearly regulated in most countries (Falconer et al., 1999). The WHO recommended multiple guidelines to evaluate the risk of MC exposure in recreational water; above

100,000 cyanobacterial cells/ml, 20 ppb MC-LR, or 50 ppb Chlorophyll-a are considered as having a high probability of acute health effects (Table 1.3). Several US states with frequent bloom concerns have developed different guidance and action levels for recreational water use (Table 1.3). Since December 2016, USEPA started collecting comments for draft guidelines of MC levels in recreational water, which considers 4 ppb of MC as the threshold for taking action. This draft will soon be finalized to provide more comprehensive guidelines for the public.

Table 1.2 Guidelines of microcystins in drinking water in different countries (modified from Carmichael and Boyer 2016). Country Cyanotoxins Guideline value

Brazil, China, Finland, France, Germany, Italy, Japan, Korea, Norway, MC-LR 1.0 ug/L (based on WHO guideline) New Zealand, South Africa, etc.

Canada MC-LR MAC: 1.5 ug/L HAG: 0.3 ug/L for 0–6 yr old; MC-LR 1.6 ug/L for school age to adult United States of America HAG: 0.7 ug/L for 0–6 yr old;

11 Cylindrosper-mopsin 3.0 ug/L for school age to adult MCs PGV: 1.3 ug/L MC-LR equivalents Australia Saxitoxins HAV: 3 μg/L Anatoxin-a 6 μg/L New Zealand Cylindrosper-mopsin PMAV: 1 μg/L Saxitoxins PMAV: 3 μg/L Cylindrosper-mopsin 15 ug/L Brazil Saxitoxins 3 ug/L MAC: maximum concentration; HAG: Health advisory guideline; PGV: provisional guidance value; HAV: health alert level; PMAV: provisional maximum value.

Table 1.3 Guidelines for recreational water (agricultural water) (modified from US EPA 2017). Organization/State Guidance/Action Level Recommended Action Cyanobacteria >100,000 cells/ml; MC-LR > 20 WHO μg/L; High probability of acute health risk. Cholorophyll-a > 50 μg/L.

Trigger: prompts increased monitoring and the Trigger: MCs= 0.8 μg/L; placement of a caution sign stating that people CA Warning: MCs= 6 μg/L; should stay away from scum and pets and Danger: MCs = 20 μg/L livestock should be kept away from the water and scum.

14 µg/L MC-LR and ≥ 70,000 cells/mL for MA Avoid contact with water. 12 cyanobacteria Swimming and wading: not recommended; not 6 µg/L MC-LR swallow water; avoid surface scum. OH Recommend the public avoid all contact with the 20 µg/L MC-LR. water. Visible scum and cell count or toxicity Toxigenic species >100,000 cells/ml OR Avoid contact with water. Microcystis or Planktothrix > 40,000 cells/mL MCs >10µg/L

1.2.3 Exposure routes of microcystins for humans

There are multiple exposure routes to MC intoxication through environments, drinking water, food, and dietary supplements. MCs have been detected in 39% of lakes within the US, and there is a significant increase of microcystins in aquatic environments

(EPA, 2016c), indicating the recreational or occupational exposure risk of MC is continuously growing. In August 2014, the City of Toledo, Ohio, found MCs in the drinking water supply affecting approximately 500,000 residents, which demonstrated the vulnerabilities of finished drinking water to cyanotoxins during bloom periods.

However, the MC exposure through food was underestimated. The bioaccumulation of MCs in crops (through contaminated irrigation water) and seafood (through contaminated aquatic water use) has been demonstrated(Lee et al., 2017). Common sense suggests that the bioaccumulation of cyanotoxins in agricultural plants is positively correlated to exposure time and concentration. Table 1.4 summarizes several recent studies associated with MC accumulation in food via pre-exposure to MCs. The concentration of MCs in food supplies ranged from non-detected to 158.35 μg/kg fw

(fresh weight). Most of these studies emphasized the high risk of MCs for human health and indicated the necessity of regulation to ensure food safety. Furthermore, the residual

MC-LR in soil can persist for a long time, which leads to the toxin accumulated into next cultivation (Lee et al., 2016; Machado et al., 2016b).

Another exposure route, algal dietary supplements, has also been investigated. The algal supplements are metabolites from non-toxic species cultured in natural water, such as Aphanizomenon flos-aquae, a common algal dietary supplement resource aimed to

“elevate mood, increase energy, and alleviate attention deficit hyperactivity disorder”

(Yakes et al., 2015). However, MC contamination seems to be inevitable since the toxic algae species usually co-occur in such environments (Yakes et al., 2015). Parker et al.

(2015) reported that their supplement sample contained 0.18–1.87 μg/g (MC-LR equivalent) MCs, including LR, LA, and LY. Roy-Lachapelle et al. (2017) tested 18 commercial dietary supplements on the market, 8 of which contained cyanotoxins at levels above the tolerable daily intake (TDI). Those results indicated that more awareness of manufacture and consumers are needed.

Interestingly, Liu et al. (2014) and Wu et al. (2016) reported that parental exposure to

MCs may result in innate hepatotoxicity, neurotoxicity, immune dysfunction, and growth inhibition of offspring in zebrafish, and similar neurotoxicity has been found in rat pups whose mothers were exposed to MC-LR (Zhao et al., 2015). These findings indicate that intergenerational inheritance may be another potential MC exposure route for newborns.

Table 1.4 Bioaccumulation and physiological changes of crops and seafood exposed to MCs Dura MC content Accumulation levels Type MC source tion Physiological changes References (ppb) in edible portion (d) Lycopersicon Pure MC-LR 10.83 ± 0.94 ug/kg fw Decrease of the capacity of (Gutiérrez- Esculentum M. aeruginosa 100 14 plants to synthesize ATP and Praena et al., 10.52 ± 6.48 ug/kg fw (Tomato) crude extract to perform photosynthesis 2013) Solanum intra- and lycopersicum extracellular 1.16 ug/kg dw (Romero- Lake Amatitlán

(Tomato) 1931 and 90 N/A N/A Oliva et al., 1

5 in Guatemala

µg/L, 2014) 1.03 ug/kg dw respectively Capsicum Increased of Lipid annuum Not peroxidation levels and (Drobac et al., (Pepper) MC solution 90 118 μg/kg dw mentioned flavonoids content, decrease 2017) of antioxidant capacity

(continued)

Table 1.4 Bioaccumulation and physiological changes of crops and seafood exposed to MCs (Cont'd) Accumulatio Dura MC content n levels in Type MC source tion Physiological changes References (ppb) edible (d) portion Taihu Lake in (Chen et al., Not mentioned N/A 3.19 ug/kg Inhibition of root development China 2012) Oryza sativa 4.9, 7.1, 12.6, Inhibition of growth and (Rice) 1, 100, 1000 (Liang et al., MC solution 7 21.2 μg/kg photosynthesis, lower protein and and 3000 2016)

dw amylose content 16

Daucus M. aeruginosa Decrease of root growth, ascorbic 28- 5.23 ± 0.47 (Machado et carota aqueous 10/ 50 acid content, changes of mineral 32 ug/kg fw al., 2017) (carrot) extracts contents Petroselinum M. aeruginosa crispum L. crude extract (parsley) (Pereira et al., 0.1, 0.5, 1 10 Not detected Not significantly difference Coriandrum Microcystis. 2017) sativum L aeruginosa (Coriander) crude extract

(continued)

Table 1.4 Bioaccumulation and physiological changes of crops and seafood exposed to MCs (Cont'd) Dur MC Accumulation atio Type MC source content levels in edible Physiological changes References n (ppb) portion (d) 10 times (Cordeiro-Araújo Pure MC-LR 5, 10 7 N/A accumulation et al., 2016) Lactuca sativa L Microcystis. Increased net photosynthetic (Bittencourt- (lettuce) 0.65, 2.5, 39.31-158.35 aeruginosa crude 15 rate, and activity of Oliveira et al., 6.5, 13.0 ug/kg fw

extract antioxidant enzymes 2016) 17

Detoxification and Corbiculaleana cyanobacterial crude 3.41 ± 0.63 μg/g antioxidant enzymes in (Pham et al., P. (clam) 400 10 extract dw different organs responded 2016)

differently H. molitrix 10.12 - 40.98 (Silver carp) Anzali wetland in 0.18 to ug/kg fw (Rezaitabar et N/A N/A E. lucius Iran 3.02 μg/L 15.18 - 35.1 al., 2017) (northern pike) ug/kg fw

(continued)

Table 1.4 Bioaccumulation and physiological changes of crops and seafood exposed to MCs (Cont'd) MC content Durati Accumulation levels Physiological Type MC source References (ppb) on (d) in edible portion changes C.carpio (common 0.37 ± 0.07 - 8.87 ± carp) 3.6 ug/kg dw C.gibelio (Prussian 0.69 ± 0.26 - 3.45 ± Lake Eğirdir in 20.5 (Gurbuz et al., carp) 1.1 ug/kg dw Turkey (maximum) 2016) A.boyeri (Big-scale 0.70 ± 0.1 to 17.54 ±

sand smelt) 7.4 ug/kg dw 1

8 N/A

N/A P.pekinensis (white 79.5 ± 56.6 ug/kg dw amur bream) C. ectenes (Japanese Lake Chaohu in Not 70.6 ± 5.9 ug/kg dw (Jiang et al., grenadier anchovy) China mentioned 2017) C. auratus (gold fish) 81.7 ± 104.2 ug/kg dw H. molitrix (Silver 90.9 ± 146.9 ug/kg dw carp)

1.2.4 Effects of MCs on agriculture

Machado et al. (2016, see also Table 1.4) summarized the effects of MCs on the development, biochemistry, and morphology of crops, which are highly dependent on the concentration of cyanotoxins, exposure time, plant type and exposure stage, and toxin type. More specifically, most ecologically relevant concentrations do not show toxic effects and even promote productivity and yields, though detrimental effects were still observed at high concentrations of MC contamination (Figure 1.4). Exposure time showed a positive correlation with inhibition. Some plant species showed higher tolerance to cyanotoxins, and overall were more sensitive to MCs at the seedling stage than the developmental stage, while plants at the maturation stage were relatively inert.

Finally, the toxin types applied by some cyanotoxins may pose synergistic effects in inhibiting the development of plants. Since the mechanisms of MCs in plants are still unclear, it is important to use the correct biomarkers to study this problem.

Figure 1.4 The effects of MC-LR and CYN on plant growth, development and biochemical processes and human health risks (from Machado et al. 2016)

1.2.5 Quantification of microcystins

Many methods have been developed to measure MCs. High performance liquid chromatography (HPLC) or mass spectrometry (LCMS) have both been applied, and are relatively accurate for quantifying each congener of MCs. However, the obvious disadvantage is that not all standards for MC variants are commercially available, and it is time-consuming and expensive to quantify MCs in natural environments where they are generally released as mixtures.

Currently, ADDA-based enzyme-linked immune sorbent assays (ELISA) and protein phosphatase inhibition assays (PPIA) are available in the market to rapidly measure the quantity and toxicity of MCs (congener-independent detection) (EPA, 2015

& 2016). According to the manufacturer’s user manual (Abraxis,), the principle of the

ELISA kit is that ADDA structures from microcystins or nodularins compete with the analogues of MCs immobilized on the plate, to bind added antibodies that generate color reaction (Figure 1.5). Therefore, the disadvantage of this method is that it cannot verify specific congeners and the structural configuration (cyclic or linearized) of MCs. Some researchers argue that ELISA results may generate false-positives if only an ADDA structure is present. The PPIA, by measuring the inhibition of phosphatase activity

(manufacturer’s manual, Zeu-Inmunotec), provides direct toxicity information on MCs and nodularins, expressed as an MC-LR equivalent. However, the presence of other phosphatase inhibitors (e.g., okadaic acid) in samples may lead to inaccurate readings.

Therefore, it is more accurate to choose methods that fit the reality, instead of pursuing a certain method without considering the real issues.

Figure 1.5 Major principle of ADDA-based ELISA (modified from https://www.lsbio.com/elisakits/mouse-p2rx7- p2x7-elisa-kit-sandwich-elisa-custom-ls-f16542/16542)

Quantitative PCR (qPCR) has also been applied to measure the MC-synthetase genes

(mcy) as an indicator of MC concentration. The mcy cluster consists of nine genes involved in the biosynthesis of MCs. Although Beversdorf et al. (2015) argued that the abundance of mcyA and mcyE were not good indicators of MC level in four lakes located in Wisconsin, these methods are still widely used due to good correlations with MC concentrations in other locations (DOI & USGS, 2013; Dong et al., 2016; Francy et al.,

2016).

1.2.6 Decontamination of MCs

Since MCs are very chemically stable, we urgently need to develop effective MC decontamination strategies to ensure water and food safety. Ultraviolet (UV) light, a traditional and widely used disinfection method, can degrade MCs when high dose irradiation is applied. The ADDA structures of MCs were isomerized at conjugated diene

[from (4E), (6E) to (4E), 6(Z) or 4(Z), 6(E)] under UV light, and no toxic byproducts 22 were detected (Tsuji et al., 1995). However, the degradation effectiveness is not only dependent on the UV dose, but easily affected by turbidity and the presence of organic matter. Therefore, UV is usually amended with other oxidants (e.g. hydroperoxide) (Liu et al., 2016a; Qiao et al., 2005; Thirumavalavan et al., 2012). UV at different wavelengths (e.g., 185nm, 254nm, 265nm, 300-400 nm, etc.) can degrade MCs (Liu et al., 2016a& b; Welker & Steinberg, 2000), but the effectiveness may differ by UV type

(UV-C> UV-B> UV-A), since the shorter wavelengths have higher energy (Eskandarian et al., 2016).

Titanium dioxide (TiO2), a widely-used photo-catalyst and food additive (Weir et al.,

2012), became popular to enhance the effects of UV light. The mechanisms of its photocatalyst are summarized in Figure 1.6. The electrons in the valence band are excited by energy from photons to the conduction band, therefore forming a positive hole and a free electron (Antoniou et al., 2009; Fagan et al., 2016; Senogles et al., 2001), which further oxidizes water or reduces oxygen to form hydroxyl radicals or superoxide radical, to attack the conjugated diene of ADDA. Non-toxic byproducts were clearly identified by

Liu et al. (2003). However, since excessive ingestion of TiO2 may cause health concerns

(Bettini et al., 2017; Golja et al., 2017; Heringa et al., 2016; Kloet et al., 2016), direct contact should be avoided during the MC treatment. Chapter 2 describes how UV combined with a reusable coating of TiO2 can effectively degrade MCs.

Figure 1.6 Schematic diagram of photocatalytic mechanism of TiO2 (from Murakami and Fujishima 2010).

Plasma, the fourth stage of matter (as opposed to solids, liquids, and gases), has been widely applied for pollutant decontamination and pathogen disinfection (Basaran et al.,

2008; Min et al., 2012; Niemira, 2012; Perni et al., 2008a). Typically, gas was ionized to plasma by different sources (e.g., heat, electricity, laser light, etc.) to produce a variety of reactive species that inactivate pathogens or decontaminate the pollutants (Patil et al.,

2016; Ziuzina & Misra, 2016). Since cold (non-thermal) plasma can maintain relatively low temperatures of targets with little thermal damage, it has been especially attractive in the food industry recently (Han et al., 2016; Misra et al., 2016; Schnabel et al., 2015).

However, not much research has been done on MC decontamination using cold plasma. 24

Zhang et al. (2012 &2014) have reported that glow discharge plasma is effective to remove MC-LR. In Chapter 2, a new cold plasma method is evaluated for treating MCs.

Chemical oxidants can remove MCs as well, and chlorine is the most convenient and commonly applied. Its effectiveness may be affected by the chlorine dose, pH of the matrix, and the presence of other organic matter (Acero et al., 2005; Chu et al., 2017). A major concern about chlorine treatment is the generation of toxic byproducts during MC chlorination, such as trihalomethanes (Tsuji et al., 1997; Zhang et al., 2016; Zong et al.,

2015). Other oxidants such as ozone, permanganate, and other chlorides may show different effectiveness due to their oxidative abilities (Rodríguez et al., 2007).

The biodegradation of MCs has attracted more attention recently. The mlr gene cluster is responsible for MC degradation via linearization by MlrA, and breakdown by

MlrB and MlrC. MlrD serves as a transporter located on the cell membrane to facilitate

MC uptake and metabolite excretion (Figure 1.7) (Dziga et al., 2013; Eleuterio & Batista,

2010; Kormas & Lymperopoulou, 2013). Many studies have reported successful isolations of MC degradation bacteria from various environments; however, biodegradation may be unfeasible for industries due to the long treatment time (Eleuterio

& Batista, 2010; Manage et al., 2009; Phujomjai et al., 2016; Yang et al., 2014).

Figure 1.7 Pathway of MC biodegradation (from Dziga et al. 2013).

Previously, researchers tended to use a single MC congener or artificial MC mixture to assess the efficiency of MC treatments. However, natural MCs are released as a mixture of many MC variants and other metabolites of cyanobacteria, which show 26 higher toxicity due to potential synergistic effects (Esterhuizen-Londt et al., 2016;

Olivares Rubio et al., 2015; Rymuszka & Sierosławska, 2013). These findings demonstrate that it is necessary to develop effective methods to remove natural MC extracts. In addition, there is a gap in evaluating MC degradation in contaminated food using the methods mentioned above. Food consists of a complex matrix including carbohydrates, lipids, proteins, minerals, and other components, which may affect the effectiveness of the MC removal. Meanwhile, maintaining high food quality should be considered when choosing appropriate treatments.

1.3 Health concerns from other cyanotoxins

Besides MCs, other cyanotoxins threaten food and water safety as well (Table

1.5). Nodularins are hepatotoxins with an ADDA structure, and therefore show similar effects of MCs. Saxitoxins, anatoxins, and anatoxin-a(s) are neurotoxins affecting the neural signaling pathway, and can lead to death. Cylindrospermopsin leads to multiorgan damage due to its toxicity against DNA. The EPA recommended limiting cylindrospermopsin in drinking water to 0.7 ppb and 3 ppb for pre-school children and adults, respectively (Table 2). Several states with frequent bloom concerns developed their own guidelines for other cyanotoxins (details can be found at the US EPA website).

Similar to MCs, these cyanotoxins may also accumulate into food and dietary supplements via contaminated water (Ibelings & Chorus, 2007; Lee et al., 2017; Pawlik-

Skowrońska et al., 2013; Roy-Lachapelle et al., 2017; Saqrane & Oudra, 2009; Temsah et al., 2016), which underscores the importance of establishing guidelines for agricultural water quality and food quality in terms of cyanotoxin contaminations.

Table 1.5 Toxicity and health effects of other cyanotoxins (modified from Wiegand and Pflugmacher 2005; Lee et al. 2017). Cyanotoxins Classification Mechanisms Health effects Inhibition of protein Nodularins Hepatotoxins phosphatases (PP1 and Similar to MCs. PP2A). Numbness, Locking the Na+-channels respiratory Saxitoxins Neurotoxin close in neural cells. paralysis to death, etc. Irreversible binding to the nicotinic acetylcholine Cerebral anoxia, Anatoxins Neurotoxin receptors causing the Na+- suffocation. channels locked open. Inhibition of Ach-esterase Anatoxin-a(s) Neurotoxin Respiration failure. activity. Multi-organ Irreversible inhibitor of damages, Cylindrospermo protein biosynthesis, Cytotoxin mutagenic and psin cytogenetic damage on potential DNA. carcinogenic. Fever, septic shock Lipopolysacchar Irritation to exposed Irritant toxin syndrome, ide (lipid A) tissues. inflammation,

2. Cyanophages-cyanobacteria interplay

2.1 Bacteriophage-bacteria interaction

2.1.1 Introduction of virus with the emphasis of bacteriophage

Since the first discovery of a unique particle that can infect tobacco (the tobacco mosaic virus) in the 1890s, viruses have come to the public’s attention. Given their special structures and life cycles, scientists have long debated viruses’ positon in the biological spectrum. It is known that viruses cannot metabolize and reproduce independently from hosts, but after invading host cells, they can take control of their life processes and produce more new viruses (Brown, 2001). Therefore, viruses are more likely to be considered as “active/inactive infectious particles”, rather than living/dead organisms, since they must parasitize their hosts intracellularly (Talaro & Chess, 2015).

Generally, viruses consist of genetic material (DNA or RNA) and capsid (outer protein shell), ranging in size from 20 nm to several hundred nanometers (mimivirus, the largest virus known so far), and some animal viruses contain envelope outside their capsids

(Ekeberg et al., 2016).

Bacteriophages (simplified as phages) are viruses that infect specific bacteria, and are considered a natural predator of microbes. They are ubiquitous in various environments from ocean to freshwater and from soil to desert, and are ten times more abundant (~107/ml) than their host bacteria (~106/ml) (Stern & Sorek, 2012). Most phages contain double-strand DNA (dsDNA) packed in a polyhedral head (capsid), as well as a helical tail (fibers) (Figure 1.8).

Figure 1.8 Diagram of bacteriophage (as example of T4 Escherichia coli phage, from Talaro and Chess 2015).

As Figure 1.9 shows, phage infection includes adsorption, penetration, replication, assembly, maturation, and releasing (Talaro & Chess, 2015). Adsorption of phages into host is mediated by the specific receptors on the bacteria’s surface that exactly fit the phages. Bacteria lacking the requisite receptors are resistant to that phage. The nucleic acid is then injected into the bacterial cell (penetration), following the replication of phage nucleic acids and production of viral parts by controlling the host’s gene expressions. The free units of phages are spontaneously assembled and released as mature bacteriophages, with the lysis of the host cell. However, instead of lysing the hosts, some of the bacteriophages recombine their DNA into the host’s genome without immediate replication. The viral DNA is carried by bacterial chromosomes and replicated with the hosts’ reproduction, until it is reactivated by external factors (induction). This stage is called lysogeny, and since this type of phage is less lethal to the host cells, they are described as “temperate” phages.

Figure 1.9 Life cycle of bacteriophages (from Talaro and Chess 2015).

The advantages of lysogeny are explained by Mann (2003): 1) exchanging the genetic material without killing the host expands the mobility of viral DNA; 2) maintaining the host’s abundance ensures its own replication; and 3) infected cells are immune to further infection with homo-immune phages, ensuring the selection of non- lysogenic hosts.

2.1.2 Co-evolution of bacteriophages and their hosts: an arms-race

Importantly, bacteriophages drive the evolution of their hosts, thereby shaping entire microbial communities. Firstly, the specificity of bacteriophages can maintain the biodiversity of an ecosystem. When a specific strain becomes dominant, its phages can rapidly expand and control the population, and leave space for other living microbes.

Therefore, the internal community structure is always in a diverse, dynamic equilibrium.

By injecting their genetic material into host cells, bacteriophages, especially the temperate phages, also provide more opportunities for gene exchange (transduction).

Phages with a broad range of hosts serve as important gene carriers between different microbes or different ecosystems, which accelerates the gene transfer rate (Rohwer et al.,

2009). The acquisition of viral genes may also favor the evolution of pathogens (Table

1.6 Bacterial virulence properties altered by bacteriophages (from Wagner and Waldor

2002).). Pathogenesis acquired from phage-encoded genes enhances the pathogens’ abilities to infect humans, which facilitates its spreading (Wagner & Waldor, 2002).

Bacterial hosts also developed protective mechanisms from phage infections, which may promote co-evolution with their phages (Stern & Sorek, 2012). In addition, the high mutation rate of viruses is a major force to improve the diversity of genetic information, as well as to provide new selective pressure against their hosts. The gene interaction between temperate phages in the same host may show even more evolutionary significance, since bacteria can harbor multiple prophages at the same time (Casjens,

2005). The recombination of host-virus, host-host, and even virus-virus genes then further impacts the evolutionary direction of the biosphere.

Table 1.6 Bacterial virulence properties altered by bacteriophages (from Wagner and Waldor 2002). Bacterial Example of human Mechanism properties altered pathogens Colonization/ The λ-encoded gene promotes adhesion to E. coli adhesion buccal epithelial cells. Phage Gifsy-2 encodes SodC, a Resistance to superoxide dismutase. S. enterica serum/phagocytes Phages encode enzymes that alter the O antigen. Exotoxin C. botulinum Botulinum toxin is phage encoded. production Generalized transduction contributes to Susceptibility to S. aureus horizontal transmission of gram-positive antibiotic antibiotic-resistance genes.

However, the genomic information of most bacteriophages, especially the prophages, is unexplored, and the details of host-phage interactions are poorly understood. With the development of whole genome sequencing and metagenomic analysis, it is expected that more mechanisms of phage infection, interaction, and evolution can be described.

2.1.3 Biocontrol of foodborne pathogens using bacteriophages

Driven by the increasing foodborne diseases globally, the effective controlling of pathogens present are required to ensure the food safety (Figure 1.10, CDC, 2016; WHO,

2010). However, traditional treatments usually cost high energy (e.g. electricity or water) and may impair the food quality (e.g. heat) and public perception (e.g. Gamma irradiation). With the call of “natural” food from consumers, bacteriophages become a good choice to reduce or eliminate the presence of pathogens in food. The advantages of

33 bacteriophages as biocontrol strategies are obvious: 1) specific infections against certain pathogens, but no adverse effects on mammal or plant cells; 2) generally being digested as normal dietary components; 3) no significant effects on gut microbes; 4) relative stability during long term storage (Goodridge & Bisha, 2011; Kazi & Annapure, 2016).

Figure 1.10 WHO estimated global foodborne disease burden (from WHO, 2010).

Bacteriophages are used in both pre- and post-harvest. The hide spray of phage cocktail in animal farms was approved by USDA for reducing E. coli O157:H7 shedding before slaughter (USDA, 2014). Other research also tested the effectiveness of oral administration of bacteriophages on mitigating the E. coli O157:H7 shedding, but the viability of those phages after digestion seems to be the major concern (Rozema et al.,

2009; Sheng et al., 2006). Therefore, the delivery of bacteriophages in animals is critical for its application as biotherapy. Meanwhile, EPA approved the use of bacteriophages as

34 pesticides in tomatoes and peppers (EPA, 2016a; EPA Office of Pesticide Programs,

2012). The effectiveness of phage-pesticides may be affected by varying environmental factors.

Application of bacteriophages in food are becoming popular now. Desirable bacteriophages should be fully sequenced, strictly lytic against with host range, do not carry genes encoding virulence and allergic proteins, and be lack of transduction activities (Goodridge & Bisha, 2011). Phage cocktail (mixture of several phages) is used to improve the effectiveness against multiple pathogenic strain, as well as to lower the risk of resistance (Kazi & Annapure, 2016). Generally, the proper dose of phage cocktail can reduce 2-4 log of targeted pathogens alone and higher efficacy can be achieved by combing with other strategies (e.g. nisin) (Tabla et al., 2012; Wang et al., 2017). Several commercial bacteriophage products targeting pathogenic Listeria, Salmonella and E. coli have been approved by the FDA in various foods (details can be found at: http://www.intralytix.com/index.php?page=prod and http://www.phageguard.com). The complexity of food matrix may be the major hurdle of wide applications: semi-solid/solid food reduces the diffusion of bacteriophages to encounter their hosts, which may present at low concentration (Hagens & Loessner, 2010; Kazi & Annapure, 2016); bacteriophages may be inactivated or less effective to infect hosts by unfavorable pH or storage temperature (Pometto & Demirci, 2015; Pulido et al., 2015).

To conclude, bacteriophages are promising as biocontrol of pathogens in various food types, but more studies are needed in development of suitable phages; improvement of efficacy in various conditions; optimization of delivery in alive organisms or food; combination with other controlling strategies.

2.2 Cyanophages

Cyanophages refers to a collective of phages targeting cyanobacteria, which has been found to be abundant in various aquatic ecosystems. Typically, cyanophages are double- strand DNA tailed-phages (Clokie & Mann, 2006; Xia et al., 2013). Majority of cyanophage studies have focused on oceans. Most marine cyanophages affecting

Synechococcus and Prochlorococcus belong to three viral families: Myoviridae, which have long straight contractile tails; Podoviridae, with short non-contractile tails or no obvious tails; and Siphoviridae, with long flexible non-contractile tails (Fokine &

Rossmann, 2014). Synechococcus is classified as marine cluster (MC) A with the photosynthetic complex, phycobilisome, and Prochlorococcus is classified as MC-B with phycocyanin as its major photosynthesis pigment (Lu et al., 2001), so their cyanophages are classified by the hosts. Marine cyanophages tend to have a broad host range that can infect multiple strains of its host and even be cross-infectious (Jia, 2009).

Freshwater cyanophages, however, are poorly described compared with the marine cyanophages. The first cyanophage LPP-1 belonging to Podoviridae was isolated in 1963 (Safferman & Morris, 1963& 1964), and subsequently other cyanophages were isolated in various freshwater systems (Table 1.7 Examples of freshwater cyanophages

(from Xia et al. 2013).). Overall, Podoviridae and Siphoviridae are more commonly found in fresh water compared to Myoviridae (Xia et al., 2013).

Since cyanobacteria are a major producer on the earth, cyanophages are thereby further related to the primary production in food web besides the functions mentioned in

2.1 (Mann, 2003b). Some researchers proposed that manipulation of hosts’ photosynthesis may facilitate phage infection (Figure 1.11, Bailey et al. 2004; Clokie and

Mann 2006; Rohwer et al. 2009; Xia et al. 2013). The psb gene present in marine cyanophages protects the photosynthetic capacity in host cells from excessive light stress, to ensure the production of energy for viral replication. Dreher et al. (2011) demonstrated the first presence of the psb gene in freshwater cyanophages. Ma-LMM01, an unique freshwater cyanophage that does not share the psb gene with other marine cyanophages, was interestingly characterized by Yoshida et al. (2006). This cyanophage contained a phycobilisome (the major photosynthesis complex) degradation gene, nblA, to direct the host’s photosynthesis and produce free amino acids (Van Wichelen et al., 2016).

Freshwater tailless cyanophages PaV-LD and Myoviridae MaMV-DC also carried the nblA gene obtained from their hosts, which negatively affected the phycocyanin contents

(Gao et al., 2012; Ou et al., 2015). Puxty et al. (2016) mentioned that marine cyanophages redirect photosynthesis by limiting the CO2 fixation during the early infection stage to save energy for viral development, which Kaplan (2016) described as

“starving hosts.” Recently, Roitman et al. (2017) found a gene encoding lipid-desaturases in marine cyanophages, which may improve the fluidity of host cell membranes and further increase the photosynthesis of host cells. All these changes in cyanobacteria photosynthesis by cyanophages may contribute to the up- or down-regulation of primary productions, so the cyanophages are critical to maintaining the global carbon cycle.

Table 1.7 Examples of freshwater cyanophages (from Xia et al. 2013). Family Phage species Host Myoviridae Ma-LMM01 Microcystis aeruginosa Siphoviridae SM-2 Synechococcus elongates, Microcystis aeruginosa Podoviridae LPP-1 Lyngbya, Plectonema, Phormidium Unassigned (Tailless) PaV-LD Planktothrix agardhii 37

Figure 1.11 Model of the metabolic pathways regulated by cyanophage host-like genes in infected cyanobacterial cells. Cyanophage host-like genes are marked in red. (From Gao et al. 2016)

* TM, thylakoid membrane; TL, thylakoid lumen; CM, cell membrane; PSI, photosystem I; PSII, photosystem II; PPP, pentose phosphate pathway; CBB,

Calvin–Benson–Bassham; HLIP, high light-induced protein; PQ, plastoquinone (oxidized); PQH2, plastoquinone (reduced); PTOX, plastoquinol terminal oxidase; Cyt b6f, cytochrome b6f complex; PCred, plastocyanin (reduced); PCox, plastocyanin (oxidized); FNR, ferredoxin-NADP reductase; Fd, ferredoxin; G-6-P, glucose-6-phosphate; R-5-P, ribose-5-phosphate.

In addition, with a broader range of hosts than general bacteriophages, cyanophages play a more important role in the evolution of their co-existing hosts for the past billion- year arms race (Van Wichelen et al., 2016; Xia et al., 2013). For marine cyanophages, the movement of ocean currents also contributes to long-distance gene exchanges. Gao et al.

(2016) reviewed the metabolic genes of cyanophages which showed distinctive genetic 38 origins. Acquisition of cyanobacterial genes seems to be the major metabolic gene sources, which are termed as auxiliary metabolic genes (AMGs). AMGs can help cyanophages maintain their fitness during infection by temporarily improving the host metabolites (Figure 1.11), to adapt to extreme environments and to recruit essential material for its own replication (Gao et al., 2016; Kaplan, 2016; Roitman et al., 2017).

For example, by encoding the high light-inducible protein (HLIP) the hli gene in cyanophages can protect the host from high light stress to ensure its own replication

(Clokie & Mann, 2006). Therefore, the active gene exchanges between cyanophages and hosts not only improve genetic diversity, but also shape cyanophages to be highly adaptive in various environments, therefore broadening the host ranges and habitats to ensure further genetic communications (Lindell et al., 2007; Paul & Sullivan, 2005a).

Chapter 3 describes a freshwater cyanophages isolated from Lake Erie, which contained the psbA gene and a special structural capsid protein gene. This finding provides more information on phage-host co-evolution.

Although the mortality of marine cyanobacteria directed by cyanophages is relatively low (Paul et al., 2002), Yoshida and her colleagues have confirmed that the seasonal abundance of toxic M. aeruginosa was correlated with its cyanophages’ population

(Kimura-Sakai et al., 2015; Yoshida et al., 2010), indicating the importance of freshwater cyanophages in bloom formation. Lysogenic cyanophages, however, tend to improve the fitness of cyanobacteria by expressing viral-encoded genes, but their detailed functions remained unclear. Interestingly, cyanophages can be pseudolysogenic such that the infected host cells live as normal even though the virus is “pursuing a lytic infection.” It

39 is more like a middle stage between the lytic and lysogenic cycles, and prone to occur under nutrient-starving conditions as a strategy of persistence (Mann, 2003b).

However, because the genetic characteristics of cyanophages are limited, describing their dynamic changes is still difficult. PCR-based screening has been widely used to detect the presence or quantify the abundance of functional genes in cyanophages, such as the capsid protein gene g20 (Stoddard et al., 2007; Wilhelm et al., 2006), tail sheath protein g91 (Kimura-Sakai et al., 2015), DNA polymerase gene pol (Chen et al., 1996;

Pope et al., 2007), and photosynthesis protein D psb (Dreher et al., 2011; Honda et al.,

2014). However, the expression of these markers is highly restricted within specific groups of cyanophages. Due to the diversity of cyanophages, those genes missed most species and are under-investigated in the viral community (Adriaenssens & Cowan,

2014). More signature genes are likely to be revealed with the development of a cyanophage genome database, and metagenomic analysis may provide a more comprehensive picture of cyanophages in the environment. Meanwhile, current isolated cyanophages only showed shift of composition of cyanobacteria, instead of significant reduction, at environmental condition (Sigee et al., 1999; Van Wichelen et al., 2016) .

The close monitor after cyanophage administration is required to minimize the adverse effects on entire ecology. Therefore, it is still far from the practical application of cyanophages as biocontrol of bloom without further understanding of the cyanophage- cyanobacteria interaction cooperated with various environmental conditions.

Chapter 2: Sustainable Methods for Decontamination of Microcystin in Water using Cold Plasma and UV with Reusable TiO2 Nanoparticle Coating

1. Abstract

Microcystins (MCs) are a family of cyanotoxins and pose detrimental effects on human, animal and ecological health. Conventional water treatment processes have limited success in removing MCs without producing harmful byproducts. Therefore, there is an urgent need for cost-effective and environmentally friendly methods for treating

MCs. The objective of this study was to develop sustainable and non-chemical-based methods for controlling MCs, such as using cold plasma and ultra violet (UV) light with titanium dioxide (TiO2) coating, which can be applied for diverse scale and settings. MCs, extracted from Microcystis aeruginosa, were treated with cold plasma or UV at intensity of 1470 μW/cm2 [high] or 180 μW/cm2 [low]. To assess synergistic effects, the outside of the UV treatment chamber was coated with nanoparticles (TiO2) prior to irradiation, which can be reused for a long time. The degradation efficiency of UV was enhanced by the reusable TiO2 coating at lower intensity (70.41% [UV] vs. 79.61% [UV+TiO2], 120 min), but no significant difference was observed at higher intensity. Cold plasma removed

MCs rapidly under experimental conditions (92%, 120 min), indicating that it is a promising candidate for controlling MCs in water without generating harmful disinfection

41 byproducts. It can be also easily and practically used in household settings during emergency situations.

Keywords: Microcystis aeruginosa; cyanotoxin; cold plasma; UV; titanium dioxide; emergency preparedness

2. Introduction

Microcystins (MCs) refer to a family of cyanotoxins produced by cyanobacteria, such as Microcystins spp., Anabaena spp., Nodularia spp., and several other cyanobacterial species (Cheung et al., 2013). They are commonly found worldwide in surface waters, including the United States (US) and Europe (Funari & Testai, 2008).

MCs can cause headache, sore throat, vomiting, nausea, stomachache, diarrhea and pneumonia in acute exposure (Lévesque et al., 2014). Chronic exposure can lead to liver, kidney, and reproductive system damage and promote cancerous tumors (Dawson, 1998;

Rudolph-Böhner et al., 1994). Multiple pathways are involved in MC exposure, such as direct ingestion, inhalation of bioaerosol or dermal contact of contaminated water (Corbel et al., 2014; Funari & Testai, 2008). According to the WHO (2003a& b), MCs are safe to consume in drinking water at levels of 1 part per billion (ppb) and to exposure in recreational water at 10 ppb. United States Environmental Protection Agency (EPA,

2006) listed MCs as one of the highest priority for health risks in drinking water. New guidelines in US for MC-LR suggest a maximum acceptable concentration in drinking water of 0.3 ppb for pre-school-age-children and 1.6 ppb for school-age-children and adults (EPA, 2015a). However, traditional water treatment steps, including coagulation, sedimentation, and disinfection, have demonstrated limited effect for removing MCs 42 during heavy bloom periods (Hitzfeld et al., 2000). Furthermore, MCs are resistant to o direct UV exposure, ozone or chlorine oxidation (common steps for disinfection in drinking water treatment) and the remaining toxins may cause health risk for water users

(Chang et al., 2015; He et al., 2015; Pinho et al., 2015; Zhang et al., 2016). For example,

MCs were detected (~2.5 ppb) in the finished drinking water in Toledo, Ohio in August

2014 (EPA, 2015a).

Plenty of research has been done on MC degradation using physical, chemical and biological methods, such as UV irradiation at different wavelength (He et al., 2015; Liu et al., 2016b); chemical oxidants (Chang et al., 2014; Zhang et al., 2016), and microcystin-degrading bacteria (Phujomjai et al., 2016). However, most previous research focused on one single variant or artificial mixture of MCs, which could have different degradation dynamics and toxicity from natural MC mixture (Pestana et al.,

2015). Therefore, there is a critical need for developing more environmentally friendly and effective methods for controlling MC contamination to protect both public health and water/food safety, especially because harmful algal blooms are expected to be more frequent and intensive globally (Ho & Michalak, 2015; Rijcke et al., 2015)

In this study, we introduced two novel techniques, cold plasma and nanoparticle application with UV, to examine the effects of both methods for removing MCs in water.

Plasma is ionized gas at normal temperature with non-selective short half-life of active species, such as •OH, which can degrade pollutants rapidly without generating toxic residuals (Dhayal et al., 2006; Hickling, 1971; Perni et al., 2008b; Selcuk et al., 2008). It has been known to be effective to remove chemical contaminants in aqueous phase (e.g. wastewater treatment) (Montie & Kelly-Wintenberg, 2000; Sarangapani et al., 2016; 43

Wang & Chen, 2011), ensure food safety by inactivating the microorganisms and removing pesticide residuals, especially in fresh produce (Min et al., 2012; Misra et al.,

2014; Thirumdas et al., 2014). Barillas et al. (2015) described the capital cost of plasma was 20% less than traditional wastewater plant. It has been also reported that cold plasma can lower the energy consumption in food industries (Thirumdas et al., 2014). But this technology has not been widely applied for MC degradation. This method only requires water itself during MC treatment, therefore it can be perceived as clean and harmless to both treated objectives and operators, which makes it as a promising alternative to current water treatment methods, possibly for fresh produce application as well.

For nanoparticle application, TiO2 has been widely used in wastewater treatment and disinfection. UV illumination induces hydroxyl radicals produced from the active electron and positive hole of TiO2, which reacts with pollutants in water (Fagan et al.,

2016; Lawton & Robertson, 1999). It was reported that TiO2 showed a great photocatalytic activity to enhance the degradation rate of MC-LR with UV (Liu et al.,

2003; Pestana et al., 2015). The UV/ TiO2 wastewater treatment is effective and cost efficient, but increases by further separation of TiO2 in the effluent (Buthiyappan et al.,

2015). In this study, the effectiveness of UV with outer coated TiO2 has been evaluated to improve the efficiency of entire treatment, as well as lower the cost of separation. TiO2 coating is anticipated to be a time- and cost-effective, safe and sustainable way of MC treatment when combined with UV irradiation.

The objective of this research was to develop and evaluate sustainable and effective methods for MC treatment by cold plasma and UV with TiO2. This research describes, for the first time, the effectiveness of cold plasma and UV combined with TiO2 coating 44 for total MC (mixture of MC congeners extracted from M. aeruginosa that was isolated from Lake Erie) degradation.

3. Materials and Methods

3.1 MC preparation

MCs were obtained from Microcystis aeruginosa that was isolated from Lake Erie.

M. aeruginosa was cultured in CT medium using sonication (three cycles of five-min sonication followed by ten-min rest) with a Sonic Dismembrator (50 watts, Fisher

Scientific Model F50 with Probe, Waltham, MA, USA). Then, three cycles of freeze- thaw lysis were applied, and the cellular extract was filtered with sterile 0.45 μm pore- size membrane (MF-Millipore Membrane Filter, mixed cellulose esters, Billerica, MA,

USA) (Marion et al., 2012).

3.2 MC treatments and measurement

The MC extract was diluted with deionized (DI) water to final concentration of 10 ppb at pH 7.4, EC -17 mV (Mettler-Toledo AG, Analytical. CH-8603, Schwerzenbach,

Switzerland). A TiO2 nanopowder solution was made in Tween 20 (Boston BioProducts,

USA): isopropanol (Fisher Scientific, USA): acetic acid (Fisher Scientific, USA): TiO2

(21 nm, Aldrich Chemistry, USA) =1:45:6:1 by weight) (Choi et al., 2007), applied evenly to the outside of a glass container (tempered soda-lime glass, PYREX®) and dried in room temperature (about 2 hours) to test the synergistic effect with UV. The UV intensities were adjusted by the distance between the lamp and targets, and measured with UVC digital light meter (General Tools UV512C, Secaucus, NY) at the central point of the treated water surface. One liter of MC extract was treated with different conditions 45 of application with constant stirring: 1) UV (UV-C 254 nm lamp, G15T8, 15W, Philips,

Germany) at 1470 μW/cm2 (high intensity) for 90 min (pre-test showed that MC concentration was below the detection limit after 90 min treatment, data not shown); 2)

2 2 UV at 180 μW/cm (low intensity) for 120 min; 3) UV with TiO2 at 1470 μW/cm (high intensity) for 90 min (pre-test showed that MC concentration was below the detection

2 limit after 90 min treatment, data not shown); 4) UV with TiO2 at 180 μW/cm (low intensity) for 120 min; 5) cold plasma (Cleaz one-touch water sterilizer, CSOL-200SL,

220 V, 60 Hz, 80 W, Haan Co. Seoul, Korea) for 120 min; 6) without treatment in dark room for 120 min (control group) (Figure 2.1). Samples were taken every 15 min or 30 min. Total MC concentrations were measured with Microcystins-ADDA enzyme-linked immunosorbent assay (ELISA) kits (Abraxis, Warminster, PA, USA) in duplicates as

Ohio EPA (2015). Under the same conditions, three sets of each type of treatment were performed.

3.3 Energy per order (EEO) of each treatment

Energy per order (EEO) of each treatment was calculated as the equation below:

푃 푡×1000 38.4×푃 EEO = = 퐶푖 푉 푙푔 ( ) 푉 푘 퐶푓

Where P is the power of equipment (kW), V is the volume of water (L) and k is the rate constant calculated in first-order model.

3.4 Statistical analysis

To evaluate toxin degradation, the data were analyzed by one-way analysis of variance (ANOVA) for whole study design and t-test for UV with and without UV treatment at each intensity setting, using SPSS Statistics 22 statistical software (IBM,

Armonk, NY, USA). For the toxin degradation kinetics, each treatment data was fitted in first- and second-order reaction models.

4. Results

4.1. Synergistic Effects of TiO2 with UV on MC degradation

With high intensity (1470 μW/cm2) of UV illumination, 69.5% (10 min), 93.1%

(30 min) and 97.6% (90 min) of microcystins in the solution were degraded as the exposure time increased. With the UV and TiO2, MCs were removed by 69.4% (10 min),

90.5% (30 min) and 98.1% (90 min), respectively. No significant enhancement by TiO2 was observed under this high UV intensity condition (p>0.05) (Figure 2.2a).

With low intensity (180 μW/cm2) UV illumination, 33.5% (30 min), 61.3% (60 min), 67.5% (90 min) and 70.4% (120 min) of MCs were degraded as the exposure time increased, but the degrees of MC degradation were lower at each exposure time when compared to the high intensity (Figure 2.2b). However, TiO2 coating on the outside of the

UV treatment container showed a significant improvement in removing MCs at this UV intensity (p<0.05): 50.2%, 69.1%, 77.7% and 79.6% at each exposure time, respectively.

These results show that UV intensity is one of the major factors affecting the MCs degradation effectiveness and higher intensity could accelerate the removal of MCs.

However, at lower intensity, the addition of TiO2 significantly enhanced the MC remediation.

4.2. Effectiveness of cold plasma on MC degradation

MCs were removed by 66.2% (30 min), 80.2% (60 min), 86.8% (90 min) and

92.0% (120 min) using cold plasma (Figure 2.3). The MC degradation rate was faster 47 than low intensity UV illumination (with or without TiO2) (p<0.05). This result demonstrates that cold plasma can be a promising candidate for future treatment of MCs.

4.3 Kinetic analysis

To better understand the kinetics of MC degradation with different types of treatment, two different models, first-order-reaction and second-order-reaction, were used, where k is the rate constants for the first-order and the second-order reaction

2 kinetics; R is the corresponding correlation coefficient; t½ represents the half-life of microcystins under certain experimental conditions (Table 2.2).

Both the first-order and second-order models could approximately describe the kinetics of all the treatments (Table 2.1 and Table 2.2). It was apparent that the k of the high intensity group of UV treatment was much higher (about four times higher in first- order) than that of the low intensity group of UV treatment, which indicated that the degradation of MCs by UV irradiation was dose-dependent. After comparing UV with

UV/TiO2 at low intensity, the rate constant of latter was found to be higher and half-lives were shorter, indicating a more rapid degradation with TiO2 coating.

For cold plasma, the obtained R2 was almost the same for both models (Table

2.2). The rate constant of cold plasma was 0.0192 min-1 in first-order model and 0.0145

(ppb*min)-1 in second-order model, demonstrating a high effectiveness for removing

MCs. Compared to low-intensity UV treatment, cold plasma significantly enhanced the

MC degradation rate in both models (p<0.01).

4.4 Energy per order (EEO)

Energy cost of each treatment (in first-reaction model) is shown in Table 2.3.

Energy per order (EEO) is electrical energy (in kilowatt-hours, kWh) required to reduce the concentration of a pollutant by one order of magnitude (90%) in 1L of contaminated water (Bolton et al., 2001). The high irradiance of UV-associated treatments cost least energy, then followed by cold plasma, UV with TiO2 at low irradiance and UV alone at low irradiance. Overall, the energy cost of treatments in this study was much lower than other non-chemical based method (Sampaio et al., 2015).

5. Discussion

Previous research on MC degradation focused on specific variants of MCs, such as MC-LR or RR (Qiao et al., 2005; Xagoraraki et al., 2006; Xin et al., 2013). Qiao et al.,

(2005) demonstrated that H2O2 enhanced the degradation of MC-RR under UV illumination. In real world situations, a mixture of MC congeners (> 100 types) were found to be released into surface water (Niedermeyer, 2014), which might suggest different resistance and kinetics to those current treatments. He et al., (2015) investigated the relative susceptibility of various amino acids of MCs to UV treatment (with hydroxyl radical) and they found that MC-YR was the most susceptible and MC-LA was more resistant than MC-LR and -RR. In this study, natural extracts of MCs released from M. aeruginosa were used to mimic the toxin residuals in natural blooms, which contains multiple variants of microcystins (detected by LC/MS, data not shown). Ohio EPA

(2015) recommended ELISA as a rapid detection method recommended to quantify total microcystins regardless of the congeners, and similar level of MC concentration 49 measured by ELISA, LC-MS/MS and LC-UV was obtained (EPA, 2016b). Therefore,

ELISA was one of the best options to collectively measure the MCs.

UV, as a traditional disinfection method in water treatment plants, has been evaluated for MC degradation (Tsuji et al., 1995). As the results herein show, the intensity of UV highly affects its effectiveness. At high intensity, UV alone could rapidly degrade MCs in a short time, whereas a low intensity UV less effectively removed MCs.

However, the general requirement of UV intensity for water treatment is two order of magnitude lower than the experimental condition of high intensity. Previous studies added TiO2 directly into the water that is treated as a catalyst. TiO2 in the water may cause another contamination problem since there might be unpredictable hazards when exposed to the excessive TiO2 and using the TiO2 increases the treatment cost. In this study, the synergistic effect of TiO2 under UV irritation was tested by coating on the outer surface of the container, which improved public perception by preventing the release of the TiO2 into the treated water. In addition, this type of application maximizes the reusability of TiO2, thereby it can reduce the cost of each treatment and lower the risk of environmental contamination of these nanoparticles. Meanwhile, the coating method saved the cost for removing the nanoparticles from the treated water compared to other

TiO2 treatment method (nanoparticles were added in water). The saved cost was not reflected in the EEO calculation. As shown in the results, lower UV dose was still effective in removing MCs due to the synergistic effect of TiO2 (additional 17% (30 min) and 9% MC (120 min) removal). Therefore, TiO2 coating could shorten UV irradiation time to achieve a desired MC degradation efficiency at relatively low UV intensity. In addition, compared to Sampaio et al. (2015) (high intensity of UV (47,100 μW/cm2) with

2 TiO2 into water directly), our remediation treatment (UV at 180 μW/cm with surface coated TiO2) was still effective (constant of first-order reaction) to remove MC degradation in water (Table 2.1). Therefore, the coating method can be considered as an alternative to direct addition when removing cyanotoxins in a sustainable and environmental friendly way in various water treatment settings.

Cold plasma is a safe and rapid treatment that is applicable for various situations with different scales and materials (Zhang et al., 2012). Compared to chemical-based methods, such as chlorination, Zhang et al. (2012) demonstrated that the intermediates and end-products produced during MC degradation by plasma treatment were non-toxic.

Previous studies (Xin et al., 2013; Zhang et al., 2012) demonstrated that plasma was effective for removal of MC-LR by attacking the conjugated carbon double bonds of the

ADDA structure. Zhang et al. (2012) showed that glow discharge plasma can degrade

MC-LR with a rate constant of 38.81 ppb-1 min-1 in second-order model (Table 2.2). This study is the first to confirm the effectiveness in MC degradation while using the natural mixture of MCs derived from the toxic M. aeruginosa that was isolated from the harmful bloom in Lake Erie.

When compared with the UV+ TiO2 (low UV intensity), the cold plasma treatment showed additional 16% (30 min) and 12% (120 min) of MCs removal at the same exposure times under the study conditions. Therefore, cold plasma treatment can shorten the MC treatment time further, which will be quite useful and impactful, especially during a high bloom period and under an emergency when toxin in the water needs to be treated in a timely manner. The EEO of cold plasma was lower than UV and

UV/TiO2 at low irradiance. The treatment time can be further shortened if relatively low 51 level of MC is, present, indicating that it could serve as a practical backup for household treatment. For example, when municipal water treatment cannot meet the USEPA guideline in drinking water (e.g. Toledo water crisis in 2014), cold plasma can be immediately used to treat the MCs in tap water in affected families. In water treatment plants, large-scale application might improve effectiveness further to achieve the same degradation effects in a shorter period. Previous studies demonstrated that powerful disinfection using cold plasma in fresh produce can be achieved without generating extra heat and destruction of food quality (Pignata et al., 2017; Schnabel et al., 2015; Selcuk et al., 2008; Wang et al., 2012). Moreover, the cold plasma did not require any reagents beyond air and water, therefore it is a green technology without generation of harmful waste and unwanted byproducts. However, few studies have been conducted on investigating the feasibility of cold plasma application in MC removal in food, which is another critical exposure route to cyanotoxins (Lee et al., 2017). Combining the results from this study, it is promising to test the effectiveness of cold plasma for degrading MCs accumulated in fresh produce, fish and other food-related matrices. Therefore, more efforts should be made in MC decontamination in food in the future.

6. Conclusions

This study reports the effectiveness of total MC degradation using sustainable cold plasma and TiO2 coating. The UV irradiation combined with TiO2-coating can enhance the effectiveness of MC degradation. Our developed technique prevents the direct contact between TiO2 and water to avoid unwanted contamination. In addition, the cold plasma degraded the toxin effectively without generating harmful byproducts. The 52 cold plasma can be a promising application for both household and industry setting using different scale of equipment. In future studies, other analytical tools other than ELISA and toxicity tests are recommended to identify the potential changes in the composition of microcystins and intermediates after each type of toxin treatment. It can be also practically used in household settings during emergency situations.

7. Figures

Microcystis aeruginosa Cultured in CT medium

Microcystins

Figure 2.1 Schematic diagram of MC treatment in this study.

(a) (b)

Figure 2.2 The MC degradation rate in water treated with UV and UV/TiO2. (a): high intensity (1470 μW/cm2); (b): low intensity (180 μW/cm2) treatment. Results represent three independent sets of duplicate measurements. The average of three sets was expressed with lines.

Figure 2.3 The MC degradation rate in water treated with cold plasma. Microcystins (final concentration of 10 ppb) were treated with cold plasma (blue circle) for 120 min or put in a dark room (as a control) (green diamond) with constant stirring. Results represent three independent sets of duplicate measurements. The average of three sets was expressed with line.

8. Tables

Table 2.1 Degradation rate constants (k) and half-lives (t½) in the first- or second-order reaction kinetics in microcystin removal by UV and UV

with TiO2.

First-order (Ln(C0/C) = k1t) Treatment Reference -1 2 k1 (min ) t½ (min) R UV (Intensity:1470 μW/cm2) 0.0394 17.59 0.89 This study 2 UV with TiO2 (Intensity:1470 μW/cm ) 0.0412 16.82 0.91 This study

57 UV (Intensity:180 μW/cm2) 0.0104 66.64 0.92 This study

2 UV with TiO2 (Intensity:180 μW/cm ) 0.0129 53.73 0.93 This study

UV with TiO2 added (~182 ppb MC-LA; 47100 0.0277 25.02 0.99 Sampaio et al., 2015 μW/cm2)

Second-order (1/C-1/C0= k2t) Treatment Reference -1 2 k2 (ppb*min) t½ (min) R UV (Intensity:1470 μW/cm2) 0.0848 1.74 0.92 This study 2 UV with TiO2 (Intensity:1470 μW/cm ) 0.0913 1.58 0.95 This study UV (Intensity:180 μW/cm2) 0.0025 46.34 0.91 This study 2 UV with TiO2 (Intensity:180 μW/cm ); 0.0028 28.09 0.93 This study

Table 2.2 Degradation rate constants (k) and half-lives (t½) in the first- or second-order reaction kinetics in microcystin removal by cold plasma.

First-order (Ln(C0/C) = kt) Treatment Reference -1 2 k (min ) t½ (min) R Cold plasma 0.0192 36.10 0.95 This study

Second-order (1/C-1/C0= kt) Treatment Reference -1 2 k (ppb*min) t½ (min) R Cold plasma 0.0150 10.88 0.93 This study Glow discharge Not Zhang et al., plasma (4025.5 ppb 38.81 0.018 mentioned 2012 MC-LR)

Table 2.3 Energy per order (EEO) of each treatment. 3 Treatment EEO (kWh/m /order) Reference UV (Irradiance:1470 μW/cm2) 27.96 This study 2 UV with TiO2 (Irradiance:1470 μW/cm ) 29.85 This study UV (Irradiance:180 μW/cm2) 110.73 This study 2 UV with TiO2 (Irradiance:180 μW/cm ) 89.26 This study Sampaio et al. UV with TiO2 added 415.71 (2015) Cold plasma 59.97 This study

Chapter 3: Discovery of lytic cyanophages and their interaction mechanisms with toxic cyanobacteria: A potential biological control of harmful algal blooms

1. Abstract

Harmful cyanobacterial blooms have been a great concern for Lake Erie, across the

US and globally. Although many previous studies have focused on the effect of environmental factors on blooms, the role of cyanophages (viruses whose hosts are cyanobacteria) in the ecology of toxic blooms in lakes is poorly understood. By influencing the growth of cyanobacteria, cyanophages contribute to controlling the formation of blooms. The information on infection characteristics by cyanophage is crucial for understanding bloom dynamics, thereby potential biocontrol. The objectives of this study were to: 1) isolate and characterize lytic cyanophages from Lake Erie; and 2) examine host-cyanophage interactions using multiple tools, including atomic force microscopy (AFM).

Cyanophages were isolated from water collected from western Lake Erie and were screened for selecting best candidates that showed lytic activity. The selected cyanophage was mixed with toxin-producing Microcystis aeruginosa (host) also originating from Lake Erie, to monitor their dynamic interactions over two weeks using optical absorbance, phycocyanin and chlorophyll-a productions and quantitative PCR.

AFM, transmission electron microscope (TEM) and PCR were performed to identify the structural and genetic characters of the cyanophage.

The cyanophage was identified as Podoviridae that infects the toxic strain of M. aeruginosa. TEM showed repeating short-tailed virus in 300 nm size with novel capsid proteins. The cyanophages inhibited the growth of hosts significantly and impaired the photosynthesis by decreasing the productions of phycocyanin and chlorophyll-a.

Structural damages of the host cell membranes and attacking mechanism over time were clearly observed by AFM. psbA gene and a special capsid protein gene gp58 were identified by PCR. Whole genome sequencing is on-going to further identify the cyanophage.

This is the first study that: 1) identified the Podoviridae from Lake Erie and characterized the host-cyanophage interactions; 2) revealed the lytic cyanophage’s structure, and 3) revealed the structural changes of M. aeruginosa after cyanophage infection and potential attacking mechanism with AFM.

Key words: cyanophage, Podoviridae, Mricrocystis aeruginosa, lytic, atomic force microscopy.

2. Introduction

Viruses (mainly referring to bacteriophages in this article) are ubiquitous in various environments and are often tenfold more abundant than the host bacteria (Stern & Sorek,

2012). Cyanophages are natural predators that infect cyanobacteria as their host. Similar to other bacteriophage, cyanophage can alter metabolism and replication of its host,

60 therefore manipulating the structure of the cyanobacterial community as well as intercede in biogeochemical cycling (Mann, 2003a; Roux et al., 2016; Xia et al., 2013). In addition, the cyanophages are an important driving force in evolution. As one of the oldest and most distributed microorganisms, cyanobacteria are shaped by their phages through gene transfer over a billion-year evolution (Gao et al., 2016; Paul & Sullivan, 2005b). Most isolated freshwater cyanophages are tailed-phages belonging to Myoviridae (with a long contractile tail), Podoviridae (short non-contractile tail), or Siphoviridae (long non- contractile tail), but there are tailless phages isolated as well (Gao et al., 2012; Ou et al.,

2015; Safferman & Morris, 1964; Tucker & Pollard, 2005; Yoshida et al., 2006).

Cyanobacterial harmful algal blooms (CHABs) in freshwater not only cause a huge ecological loss, but also pose great concerns for environmental and public health.

Therefore, cyanophages are considered as a potential biological control for CHABs

(Cheung et al., 2013; Van Wichelen et al., 2016). Yoshida et al. (2008, 2010) demonstrated that the shift of toxic Microcystis aeruginosa populations may be affected by the abundance of cyanophages (Ma-LMM001-liked), indicating that cyanophage may play a critical role in bloom formation. Current PCR-based diagnoses of cyanophages were developed using structural genes in cyanophage (mostly in Myoviridae), such as the capsid protein gene g20 or tail sheath protein g91. However, the presence and abundance of other cyanophage families, such as Podoviridae and Siphoviridae, in environments are highly underestimated (Adriaenssens & Cowan, 2014). Therefore, developing more powerful genetic indicators of freshwater cyanophages is a key initial step for the high- throughput screening and precise quantification of freshwater viral communities.

For visualization, the development of microscopy with high-resolution 3D imaging, such as atomic force microscopy (AFM), enables a more accurate description of host- phage interactions. Previous studies described the application of AFM as a versatile tool to explore phage from a morphological to molecular scale (Baclayon et al., 2010;

Kuznetsov & McPherson, 2011). Specifically, the cantilever of AFM is oscillating mechanically, thus provides amplitude and phase information on the specimen. The phase image provides material difference based on the stiffness and adhesion between the cantilever tip and the surface of the specimen. It has been used for virus observation recently (Kuznetsov & McPherson, 2011; Watkins et al., 2014).

To better understand the characteristics of cyanophage in freshwater in this study, we screened water samples collected over several years from different spots in western Lake

Erie during the bloom seasons. The study is designed to: 1) isolate and characterize lytic cyanophages, both morphologically and genetically, and 2) to examine the host- cyanophage interactions using multiple tools, including AFM.

3. Material and methods

3.1 Collection, concentration, and screening of lytic cyanophage

Water samples were collected from western Lake Erie from 2013 to 2015 at different locations (Table 3. S1). Viruses in water samples were collected and concentrated using cation-coated filter methods described by Haramoto et al. (2005).

Briefly, 500 ml of water was passed through an Al3+-coated 0.45µm filters

(HAWG047S6 EMD Millipore Filter, SIGMA-ALDRICH Co., St Louis, MO, USA), and

62 viruses captured by the filter were eluted into 10 ml of 1.0 mM NaOH (pH=10.8) after rinsing with 200 ml of 0.5mM H2SO4 (pH=3.0). The eluate was neutralized with 100 μl of 50 mM H2SO4 (pH 1.0) in 10 ml of 1× Tris-EDTA buffer (pH 8.0), and concentrated by centrifugation at 3,000g for 10 min (twice) using a Centriprep™ YM-50 Filter (4310 centrifugal concentrator regenerated cellulose 50kDa NMWL, EMD Millipore, Billerica,

MA, USA).

Fifty microliters of each concentrate were inoculated into an optimized well-assay containing 100 μl of Microcystis aeruginosa culture, which was originally isolated from

Lake Erie, and incubated at room temperature with a twelve-hour light cycle for 2 days to screen for lytic cyanophages.

3.2 TEM

Ten microliter of the cyanophage propagation was mixed with 10 μl of 5% of

Glutaraldehyde to fix. Carbon/Formvar coated copper grids were glow discharged in an

PELCO easiGlow™ discharge unit. After which, a 10 ul drop of sample was incubated on the grid for 5 minutes, then incubated with a drop of 1% uranyl acetate for 30 seconds after removing excess sample. Grids were imaged in an FEI Tecnai™ G2 Biotwin TEM

(ThermoFisher Scientific, Grand Island, NY, USA) at 80 kEV and images captured using an AMT camera and software (Woburn, MA, USA).

3.3 Presence of special genes using PCR

The DNA of cyanophage concentrate was extracted using Powerviral

Environmental DNA Isolation Kit (MO BIO, San Diego, CA, USA) for further analysis.

The PCR were performed as described in Table 3.1. Primers of gp55, gp57 and gp58

(knob-like protein genes) were generated from sequences of corresponding genes (Gipson et al., 2014; Pope et al., 2007) using NCBI Primer Blast. Genes were checked by 2% agarose gel at 50V for 30 min and purified by QIAquick PCR Purification Kit (Qiagen,

Germantown, MD, USA) following manufacturer’s instruction. Sequencing was performed by BigDye® Terminator Cycle Sequencing combined with 3730 DNA

Analyzer (ThermoFisher Scientific, Grand Island, NY, USA) and identified by NCBI

BLASTn (Altschul et al., 1997). The phylogenic tree was generated using www.phylogeny.fr (Anisimova & Gascuel, 2006; Castresana, 2000; Dereeper et al.,

2008; Edgar, 2004; Guindon & Gascuel, 2003).

3.4 The effects of cyanophage infections on Microcystis aeruginosa

Cyanophage concentrates (four independent sets) or autoclaved cyanophage concentrates (as a control) were inoculated into a Microcystis aeruginosa culture (host) at a ratio of 1:100 by volume, then incubated at room temperature with a twelve-hour light cycle for 2 weeks. The optical density at 680 nm phycocyanin and chlorophyll-a concentrations were measured to monitor the dynamic changes of the host using a spectrophotometer (4001/4 Spectronic Unicam, Moyer Instruments, Inc. Tamaqua, PA,

USA) or an AquaFluor Handheld Flourometer (8000-010 Turner designs, San Jose, CA,

USA) with units of phycocyanin and in vivo chlorophyll-a (details can be found at http://www.turnerdesigns.com/t2/doc/spec-guides/998-8081.pdf) as described by Marion et al. (2012). DNA of Microcystis aeruginosa was extracted using ZR fungal/bacteria

DNA MicroPrep™ Kit (Zymo Research Corp, Irvine, CA, USA) following the manufacturer’s instruction. qPCR was performed in duplicate to quantify the host

64 concentration using mcyE primers and probes (Table 3.1). 20 μl of the PCR reaction mixture containing 2 μl extracted DNA, 0.5 μM of each primer, 0.125 Μm probe, 10 ul of TaqMan® Universal PCR Master Mix II (ThermoFisher Scientific, Grand Island, NY,

USA). CFX96 TouchTM Real-time PCR Detection System (Bio-Rad, Hercules, CA,

USA) was used.

3.5 AFM

Gelatin-coated mica was prepared as described in Allison et al. (2011). 1 ml of cyanophage propagation was centrifuged at 13,000 rpm for 15 min, and the supernatant was discarded. The pellet was mixed with 100 ul of 2.5% of Glutaraldehyde to fix overnight at 4°C, which was then applied on the gelatin-coated mica, using a pipette tip and rested for 10 min. The mica slides were washed by sterilized deionized water to remove extra propagation material and dried at aseptic atmosphere for imaging. Imaging was performed in tapping mode of a commercial AFM (MFP-3D infinity from Asylum

Research) equipped with silicon cantilever (natural frequency 70kHz, spring constant 2

N/m from Asylum Research).

3.6 UV-induced activation of the prophage

One ml of propagated phage (four independent sets) or sterilized DI water was inoculated into 300 ml of M. aeruginosa (OD680nm ~0.05) and cultured as described above. Prophage cocktail was treated with UV light with three cycles of ten-second- irradiation following 10-second-resting every day for three days. The total dose was

23.58 mJ/cm2 following the equation below:

UV dose = intensity ∗ exposure time

The effects on M. aeruginosa were measured with the same methods described above.

3.7 Statistical analysis

Data were fitted into a linear regression model that was used to describe the growth of host cell infected by cyanophages using SPSS statistic 24 (IBM).

4. Results

4.1 Screening for lytic cyanophage

Among all the samples that had been screened, one cyanophage (named as Ma-LEP below) collected at from the location A in Lake Erie (Table 3. S1) in July 2015 showed the highest lytic activities against M. aeruginosa (Figure 3. S1a). The color of the host culture turned yellow instead of green (Figure S1b).

4.2 TEM

Repeating short-tailed viruses of ~300 nm in size were observed by TEM (Figure

3.1). Interestingly, the special structure of capsids (Figure 3.1c) was similar to the knob- like proteins found in marine phage Syn5 as reported by Gipson et al. (2014), indicating a novel structure shared by cyanophages isolated in marine water and freshwater.

4.3 Presence of psbA and gp58 genes

The psbA gene was detected by PCR (Figure 3. S2a) and yielded a 582 bp gene fragment (Table 3. S2). The sequence showed high similarities compared with psbA genes found in other marine cyanophages (Figure 3.2). Primers of gp58, one of the knob- like protein genes, yielded a ~300bp-fragnent similar to the gp58 of Syn5 (Gipson et al.

2014, Figure 3.S2b, Table S3.2), which further confirmed the presence of this novel

66 protein and similar evolutionary roots of the two cyanophages. However, due to the limited reference sequences that are available, no further analysis could be done.

4.4 The effects of Ma-LEP infection on M. aeruginosa

Ma-LEP impaired the growth and photosynthesis of M. aeruginosa compared with a M. aeruginosa host using autoclaved cyanophage concentrates (control). As Figure

3.3a shows, a lower optical density (OD) at 680nm was observed compared to the control. The qPCR results (Figure 3.3b), using the mcyE gene as a proxy of microcystin production potential in M. aeruginosa population, further confirmed that the toxin productivity was down-regulated significantly at days 4, 5, and 14 (p<0.05). The M. aeruginosa in the control group may have achieved its highest environmental capacity in the experimental condition at day 5, which caused a dramatic drop at day 6. Noteworthy, the genes from cells may be stable for several days in various environments (Nielsen et al., 2007). Therefore, the remaining population of hosts might be overestimated, especially when the cyanophage lysed the M. aeruginosa. Photosynthesis is an important characteristic of cyanobacteria, differentiating themselves from other bacteria, as well as for its autotropic life (Sciuto & Moro, 2015; Shaw, 2016). Phycocyanin and chlorophyll-a are most important pigments involved in photosynthesis of M. aeruginosa and their productions greatly declined after the infection by Ma-LEP (Figure 3.3c& d).

The raw data from Figure 3.3 were fitted in linear models by SPSS statistic 24 (for mcyE gene, only data from day 0 to day 5 of both groups were fitted due to the periodic shift of populations). Table 3.2 describes the growth of pigment production rate (slope) and the correlation coefficient (R2) of each linear model. The phage-infected M.

67 aeruginosa showed significantly decrease in growth rates [measuring either OD at 680nm or the gene copy number of mcyE (p<0.01)], indicating that Ma-LEP can slow down or even prevent bloom formation. Meanwhile, production of the photosynthesis pigments, phycocyanin and chlorophyll-a, was significantly reduced after Ma-LEP infection, possibly due to the cell lysis or suppression of related genes by cyanophage.

4.5 AFM

The amplitude trace images (the left column in Figure ) provide 3D information on the cyanobacterial hosts by changing the swing of the tip; the phase trace images (the right column in Figure 3.4) show the viscosity or elasticity of the targeted objects, by reflecting the speed of tip movement due to the nature of the surface. Combining the changes from both traces, the scenario of cyanophage infection can be interpreted.

Initially, the M. aeruginosa was observed in a clear circular shape in contrast to the background (mica, in this case), indicating the intactness of host cells (Figure 3.4a).

Following cyanophage infection, certain parts of the cell membrane are susceptible to

Ma-LEP adsorption, indicating the position of receptors. At this time, the host cells were relatively intact in shape, maintaining a high contrast from the mica (Figure 3.4b).

Gradually, with replication and release of Ma-LEP, the cell tended to become irregular and broken down after a decrease of surface elasticity (Figure 3.4c& d), until they were entirely ruptured (Figure 3.4e).

4.6 UV-induced activation of the prophage

Ma-LEP also showed lysogenic activity during continuous culturing under lab conditions. However, it can be induced into lytic stage by UV irradiation at an

68 appropriate dose. Figure 3.5 shows the dynamic changes in M. aeruginosa population and photosynthesis capacity. At first, two cycles of UV irradiation did not show any effects on host growth, but after the third UV treatment, the host rapidly entered the lytic stage and host cells started being lysed. In this study, 23.58 mJ/cm2 of UV intensity was sufficient for the cyanophage to activate cell lysis. The OD at 680 nm slightly decreased in one week. In contrast, the uninfected cells showed a steady increase (Figure 3.5a).

However, qPCR results, using the mcyE gene, demonstrated that toxin productivity slightly increased, then followed a decrease at day 9 (Figure 3.5b). This may be explained by the delayed response of gene due to the stability of DNA as mentioned above. The photosynthetic pigment production, however, showed a dramatic decrease after UV treatment for both groups (Figure 3.5c& d). Ma-LEP may thus have slightly enhanced the destruction of photosynthesis by lysing the infected cells.

The raw data from Figure 3.5 (after induction, from day 5 to day 11) were fitted in linear models by SPSS statistic 24. Table 3.3 describes the growth of pigment production rate (slope) and correlation coefficient (R2) of each linear model. The phage-infected M. aeruginosa showed a significant decrease in growth rate by measuring either OD at

680nm (p<0.01), but the other three models were not statistically significant.

5. Discussion

Ma-LEP isolated from Lake Erie showed a lytic or an inducible lysogenic life cycle in toxic M. aeruginosa. The growth of the host M. aeruginosa was significantly slowed after phage infection; though the mechanisms of whereby cyanophage manipulate cyanobacterial metabolism are still unclear. However, by regulating photosynthesis

69 capacity in the host, cyanophage may take advantage of its own replication (Bailey et al.,

2004; Clokie & Mann, 2006; Rohwer et al., 2009; Xia et al., 2013). The homologue of the psb gene, originating from the cyanobacteria photosystem II core protein D1, has been identified in marine cyanophages and serves to protect host cells from light stress, which may cause the overproduction of oxidative species and damage to the photosynthetic complex. This mechanism, in turn, ensures the production of energy required by viral replication (Gao et al., 2016; Honda et al., 2014). Dreher et al. (2011) first demonstrated the presence of the psb gene in freshwater cyanophages S-CRM01, infecting Synechococcus. The presence of the psbA gene in Ma-LEP shows further evidence of this mechanism. In the lysogenic cycle, the viral psbA gene may be supplementary to host photosynthesis to provide essential energy and material for host growth. Therefore, phycocyanin and chlorophyll are increased before induction (Figure

3.5c& d, days 1-5). With a low dose of UV irradiation, expression of the viral psbA gene may be a major repairing pathway in the photosystem; however, photo-stress caused by

UV is irreversible via this mechanism, Ma-LEP starts to be lytic to save and spread its own genes. Therefore, the pigment concentration dramatically decreased in both groups due to severe damage to the photosynthesis capacity of M. aeruginosa. More evidence of quantitative gene and protein expression are needed to support this hypothesis.

Synechococcus are a major marine cyanobacteria, but also inhabit in freshwater.

Through gene transfer from host to phages in both marine water and freshwater, the psbA gene may be shared by various cyanophages and can serve as a ubiquitous indicator for screening or quantification (Adriaenssens & Cowan, 2014; Honda et al., 2014). The

70 phylogenic analysis indicated that the psbA gene, carried by Ma-LEP, showed a high similarity with marine cyanophage, which suggests a common origin or overlapped host range. Interestingly, the freshwater cyanophage Ma-LMM01 isolated in Japan did not share the psb gene with other cyanophage (Yoshida et al., 2006). However, Ma-LMM01 contains a phycobilisome (a major photosynthesis complex, especially in Synechococcus) degradation gene (nblA) to direct the host’s photosynthesis and produce free amino acids

(Van Wichelen et al., 2016), indicating a novel evolutionary route distinguishing it from

Ma-LEP. Freshwater tailless cyanophage, PaV-LD and Myoviridae MaMV-DC, also carry the nblA gene transferred from their hosts, which showed negative effects on the phycocyanin contents (Gao et al., 2012; Ou et al., 2015).

Pope et al. (2007) and Gipson et al. (2014) reported a special structure of capsid protein in cyanophage Syn5, called “knob-like protein.” Three potential genes encoding this protein were identified and sequenced, but there are no more reports of the presence of this protein in other cyanophages. This special protein may be a stabilizer for viral capsids. Unlike stabilizing proteins in other viruses “sewing” capsomeres together, the knob-like protein of Syn5 displays a distinctive diagonal positioning on the hexametric capsomeres of mature Syn5 capsid and “breaks” the symmetric structure of capsids

(Gipson et al. 2014, supplementary Figure S6). TEM images of Ma-LEP show that the capsid was decorated by special structures and then was confirmed to be homologous to

“knob-like protein” in Syn5 by presence of gp58. It can be predicted that Ma-LEP may possess similar arrangements of capsid proteins, but more details need to be described using advanced techniques. More importantly, current genetic diagnosis of cyanophage is

71 still difficult. PCR-based screening uses functional genes as an indicator of environmental viral abundance. However, the expression of most of the functional genes mentioned above is subject to a specific group or family of cyanophages. It seems to be impossible to use a single or several “universal” indicators to track highly diverse cyanophage, leading to under-estimates of this viral community (Adriaenssens & Cowan,

2014). Therefore, the investigation of more signature genes (such as gp58) can favor the detection and quantification of novel cyanophage.

AFM is an emerging technique in biological studies, including cells, viruses, and biological molecules (DNA, protein, etc.) (Almonte & Colchero, 2017; Kasas et al.,

2013; Kuznetsov et al., 2013; Rebelo et al., 2013; Whited & Park, 2014). AFM enables observation of biological specimens in fluid, which can highly maintain the bioactivity of samples and allows more vivid visualization (Almonte & Colchero, 2017). In this study, the tapping mode of AFM was used to observe the host-phage interaction. As Figure 3.4 shows, the M. aeruginosa cell showed clear structural changes due to the Ma-LEP infection. The loss of cell membrane intactness was an indicator of lysis by cyanophage, finally leading to rupture of the entire cell. Actually, tapping mode can achieve nanoscale resolution without damaging the samples (either by sample preparation or imaging steps), allowing repeated observations and flexible applications (Kuznetsov &

McPherson, 2011). Therefore, further observation of viral topology, especially the arrangement of the “knob-like protein” on Ma-LEP capsids, can take advantage of AFM tapping mode. This method also allows capture the lytic cycle in vivo, which may provide more information on host-phage interactions and mechanisms of interest.

To better understand the genetic and structural characteristics of Ma-LEP, the whole genome sequence must be performed (This is on-going, but will not be included in this thesis). By getting more information from its genome, we may predict the structure of the knob-like protein and other structural proteins contributing to the phage infection. With the development of a cyanophage genome database, it is likely that putative gene markers can be found, facilitating the rapid, high-throughput detection of cyanophage. The metagenomic analysis may provide a more comprehensive picture of cyanophages in various environments, as well as their hosts (D’Agostino et al., 2016; Mohiuddin &

Schellhorn, 2015).

6. Figures

a) b)

c) Figure 3.1 TEM images of repeating cyanophages (red arrows in a& b) with short tail (red circle in c) and special capsids (red arrows in c).

Cyanophage LEP_psbA gene

Figure 3.2 Phylogenic tree of Cyanophage LEP based on the sequence of psbA gene

a) b)

d) c)

Figure 3.3 The dynamic changes of M. aeruginosa infected by Cyanophage LEP in two weeks. a): optical density (OD) at 680nm; b) gene copy number of M. aeruginosa in log scale; c) the concentration of phycocyanin; d) the concentration of chlorophyll-a.

Amplitude trace Phase trace

Figure 3.4 Scenario of cell lysis of M. aeruginosa caused by Cyanophage LEP infection. Host lysis was shown in time series (a-e). (continued)

Figure 3.4 Scenario of cell lysis of M. aeruginosa caused by Cyanophage LEP infection (con’d).

Figure 3.5 Induction of lysogenic Cyanophage LEP by UV irradiation. Infected M. aeruginosa was irradiated by UV light at day 1,2, and 4 and following monitored in one weeks. a): Optical density (OD) at 680nm; b) gene copy number of M. aeruginosa in log scale. c) the concentration of phycocyanin; d) the concentration of chlorophyll-a. (continued)

d) Figure 3.5 Induction of lysogenic Cyanophage LEP by UV irradiation (con’d):

7. Tables

Table 3.1 PCR condition and primer sequences used in this study Genes Function Primers Condition References Capsid Denaturation at F: CAT WTC WTC CCA HTC TTC; g20 assembly 95°C for 5 min, Adriaenssens R: SWR AAA TAY TTI CCR ACR MAK GGA TC protein 35 cycles of & Cowan,

81 F: AGY GAG TTY CGC CTT AHT GT; denaturation at 2014;

g91 Tail sheath R: GRT GAY TGR CGT ACY ARR GC 95°C for Kimura-Sakai Mydoviridae Major capsid F: GTT CCT GGC ACA CCT GAA GCGT; 1 min, et al., 2015 mcp protein R: CTT ACC ATC GCT TGT GTC GGC ATC annealing at 51- F: GTG AGT GCC ATT CCT GC; 60°C for 30 Honda et al., nblA Phycobilisome R: TCT TCT TGA TGA TAG CCGC min, and 2014 degradation F: GGA GAA TTC CAA ATG GAT ACA GA TTC; extension at Ou et al., nblA protein R: GTG TAA GCT TTT AGG CGG GGT 72°C for 1 min, 2015 (continued)

Table 3.1 PCR condition and primer sequences used in this study (con’d) Photosynthesis F: TAY CCN ATY TGG GAA GC; psbA protein D1 R: TCR AGD GGG AAR TTR TG Dekel-Bird et F: CCN AAY YTN GSN CAR GT; al., 2013 pol Polymerase R: TCR TCR TGN AYR AAN GC and a final F: TGG GTG GTT ACT TCT TCC TT; extension at Podoviridae gp55 R: AAT CTC GTC GAT GTC AGA CT 72°C for 10 Knob-like F: TAC TAC CAA TGA TGC TGC TC; min. gp57 proteins on This study R: GAT CCG CCT TAT GTA GAT CC

capsid 8

2 F: CAA GGC TCT GAA CGA ATCC

gp58 R: GGT GAG AGC AGT GAT GTTG 50°C for 2 min, F: AAG CAA ACT GCT CCC GGT ATC 95°C for 10 Rantala- Microcystis Microcystin R: CAA TGG GAG CAT AAC GAG TCAA mcyE min, 40 cycles Ylinen et al., aeruginosa production Probe: 6FAM CAA TGG TTA TCG AAT TGA of 95°C for 30s, 2011 CCC CGG AGA AAT TAMARA 62°C for 1 min.

Table 3.2 The growth/ pigment production of M. aeruginosa with or without Cyanophage LEP infection as a function of time in a linear model. a OD680nm Log (gene copy number) Phycocyanin Chlorophyll-a Treatment Slope R2 Slope R2 Slope R2 Slope R2

Control 0.04 >0.99 0.28 0.82 13.10 0.93 3.45 0.84 Cyanophage 0.03** 0.99 0.13** 0.74 5.75** 0.68 2.18** 0.90 LEP

a data from day 1 to day 5 were fitted ** p<0.01

Table 3.3 The growth/ pigment production of M. aeruginosa with or without lysogenic Cyanophage LEP infection after UV irradiation as a function of time in a linear model. a OD680nm Log (gene copy number) Phycocyanin Chlorophyll-a Treatment Slope R2 Slope R2 Slope R2 Slope R2 Control 0.02 0.96 0.35 0.80 -15.44 0.94 -1.26 0.99 Cyanophage -0.01** 0.48 0.25 0.62 -18.41** 0.87 -3.15 0.83 LEP a data from day 5 to day 11 were fitted ** p<0.01

8. Supplementary Data

a) c) Figure 3.6 Color change of host culture. a):96-well assay to screen the potential cyanophage (in duplicate). One sample isolated in Crib Center showed lytic characteristic (red arrow). b): M. aeruginosa itself (left one) and infected by Cyanophage LEP (three flasks on the right). c): UV- induced prophages showed lytic activity against M. aeruginosa (right) compared with control (left).

Figure 3.7 Bands of corresponding genes on 2% argarose gel. a): psbA gene; b): gp58 gene. a) 700 bp b) 300 bp

Table 3.4 Sampling sites in Lake Erie. Sites X Y A 41.701371 -83.262634 B 41.715630 -83.454826 C 41.733299 -83.297000 D 41.742417 -83.401367 E 41.749800 -83.103683 F 41.820817 -83.184500 G 41.789467 -83.357000

Table 3.5 Gene sequences of host and Cyanophage LEP. Gene name Sequences

TCATGCTCNTGATCCACTTCTAGNNTGNAATTGTTCNATTTGTCCCTCCCAGGTTTTCTG CATCTNNTTGNNCCTTCNNCANAATTTCCCNATCGAAATTACCGGCCATGCACCTAAGA AGAAGTGCAGAGAACGGCTATTATTGAAAGAAGCGTATTGGAAGATTAAGCGTCCGAA psbA of GTAACCGTGAGCGGCAACGATATTGTAGGTTTCTTCCTCTTGACCGAATTTGTAACCGT Cyanophage AGTTCTGAGATTCGATTTCAGTGGTTTCACGCACTAAGGAAGAAGTTACTAACGAACCG TGCATTGCACTGAACAAGGAACCGCCGAACACACCAGCAACACCTAACATATGGAAGG LEP GGTGCATCAGAAAGTTATGTTCCGCTTGGAACACGCACATAAAGTTAAAGGTTCCAGA GATTCCCAAAGGCATANCATCAGAGAAGGAACCTTGTCCNATGGGGTNGATTAGGAAT ACCGCNGTTGCGNCGGATACAGGGGCAGAGTANGCTACACNNATACNGGGGGCNTNCC TAAACGGANAGANCTTCCCACTNNNACCNAGGTATCNNAAAAAACCTATAAAAAG gp58 of ACATGGCCTAATAGAGCTNAGGAATTCGCTCGGCCACATCGNCCAAAGCGATTAACAT Cyanophage AGTGATAGGAATATCCGATTCTTTGCGGAGTTCCTGACAGACACCGTAACCGTCTAATT LEP TCGGCAACATCACTGCTCTCACCAG

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