Bioprocessing of Soybean Seed-Coats for Production of Proteins & Omega

BIOPROCESSING OF SOYBEAN SEED-COATS FOR PRODUCTION OF PROTEINS & OMEGA-3 FATTY ACIDS USING PYTHIUM ISOLATES

Carren Nyambare Burkey

A Thesis

Submitted to the Graduate College of Bowling Green State University in partial fulfillment of the requirements for the degree of

MASTER OF SCIENCE

August 2020

Committee:

Paul Morris, Advisor

Kevin Neves

Vipaporn Phuntumart

Travis Worst ii ABSTRACT

Paul Morris, Advisor

Industrial processing of soybeans to produce soy oil and soy meal results in soybean seed coats as a low value waste product that is underutilized. Pythium are rapidly growing plant pathogens that secrete a large number of carbohydrate-digesting enzymes that possess the ability to breakdown the fibrous carbohydrates present in soybean seed coats to release the required carbon for their growth. We tap into this potential of oomycetes and utilize soybean seed coats as a source of carbon for the growth of Pythium under microaerobic, aerobic conditions and 4% sucrose supplementation to produce proteins and the important omega three and omega six fatty acids that can be included in fish feed as an alternative source of fish oil and proteins. Biomass produced by growing cultures on this carbohydrate source and inorganic N has a protein content exceeding 20% with a very favorable amino acid profile. Microaerobic culture conditions produce an unsaturated and saturated fatty acid mixture that mimics that of salmon and sardine oil. Aeration of samples and supplementation with 4% sucrose during the growth process significantly increases the amounts and variety of omega fatty acids produced. The results of this study suggest that soybean seed coats are a viable source of carbon for Pythium in the production of omega three and omega six fatty acids and proteins for aquaculture. iii

I dedicate this work to my son. iv ACKNOWLEDGEMENTS

I am deeply indebted to my advisor, Dr. Paul Morris, Department of Biological Sciences,

Bowling Green State University and wish to earnestly thank him for his undying guidance, invaluable advice and supervision given to me throughout this study. I am deeply grateful to my committee members Dr. Vipaporn Phuntumart, Dr. Kevin Neves and Dr. Travis Worst,

Department of Biological Sciences, Bowling Green State University for the invaluable assistance and insight provided throughout the study.

I acknowledge my mentees Robert Charles Lince, Alexandra Adrian Green, and Andrea

Cristina Cauley for their help during the study. I thank my lab mates for any support offered during the course of my research. Finally, I thank my wonderful family for their constant encouragement and motivation throughout the study. v

TABLE OF CONTENTS

Page

CHAPTER I: INTRODUCTION ...... 1

1.1 Omega Fatty Acids ...... 1

1.2 Oomycetes ...... 4

1.3 Potential of Pythium to Synthesize Omega Fatty Acids Using Various Sources

of Carbon ...... 5

1.3.1 Effects of cultural conditions in the biosynthesis of omega fatty acids

...... 7

1.4 Research Question ...... 8

1.5 Hypotheses ...... 8

1.6 Goals of the Study ...... 9

CHAPTER II: HPLC ANALYSIS OF FATTY ACIDS ...... 10

2.1 Introduction ...... 10

2.1.1 Overview of the HPLC process ...... 10

2.1.2 Derivatization of fatty acids as Naphthacyl derivatives ...... 12

2.2 Materials and Methods ...... 12

2.2.1 Raw materials ...... 12

2.2.2 Reagents and chemicals ...... 13

2.2.3 Isolation and propagation of Pythium ...... 13

2.2.4 Cultivation of Pythium isolates ...... 14

2.2.5 Optimization of growth and FA production by Pythium ...... 15

2.2.6 Fatty acid extraction ...... 16 vi

2.2.7 Saponification ...... 16

2.2.8 Goal 1: A derivatization procedure and HPLC separation method...... 17

2.2.9 Separation of fatty acid Naphthacyl esters by HPLC ...... 17

CHAPTER III: RESULTS AND DISCUSSION...... 19

3.1 Goal 2: Fatty Acids Standard Curves ...... 19

3.2 Analysis of Fatty Acid Profiles from the Isolates ...... 24

3.3 Goal 3: Optimization of Omega-3 FA Production in Pythium ...... 27

3.4 Goal 4: Effect of Sucrose and Oxygen Supplementation on FA Concentration

in the Pythium Biomass ...... 29

3.5 Quantification of Fatty Acids ...... 32

3.6 Stability of the Naphthacyl Derivatives ...... 35

3.7 Reproducibility ...... 36

3.8 Goal 5: Amino Acid Analysis ...... 36

CHAPTER IV: CONCLUSIONS AND FUTURE DIRECTIONS ...... 39

REFERENCES ...... 44 vii

LIST OF FIGURES

Figure Page

1 Fatty acid standard curves ...... 20

2 Chromatogram of a mixture of fatty acid standards ...... 23

3 Chromatogram of HPLC elution of isolate 5 (WC005) ...... 25

4 Chromatogram of HPLC elution of isolate 7 (WC007) ...... 25

5 Chromatogram overlays of the fatty acid profiles of sardine oil (fish oil) and isolate

WC007 ...... 26

6 Experimental set up during supplementation of 4% sucrose and bubbling oxygen .. 30

7 Chromatogram of HPLC elution for WC004 and WC007 under 4% sucrose and

oxygen supplementation ...... 31

8 Relative amounts of individual omega-3 fatty acids in the isolates under hypoxic

conditions ...... 34

9 Relative amounts of individual omega-3 fatty acids in the isolates under

supplementation with 4% sucrose and 500ml/min oxygen...... 34

viii

LIST OF TABLES

Table Page

1 Resolution of derivatized fatty acids using Gemini 3-micron NX-C18 110Å 250mm

column ...... 24

2 Temperature shift combinations for optimization of omega-3 fatty acid production 28

3 Analysis of fatty acids in μg/mg dry weight of cultures in flasks under hypoxic

conditions for isolate WC007...... 32

4 Analysis of fatty acids in μg/mg dry weight for isolate WC007 under 4% sucrose

supplementation...... 33

5 Analysis of fatty acids in μg/mg dry weight isolate WC007 under 4% sucrose and

oxygen supplementation ...... 33

6 Stability analysis of Naphthacyl derivatives of fatty acids ...... 35

7 Reproducibility of the derivatization technique ...... 36

8 Summary of the amino acid profile in the resulting Pythium biomass ...... 38

9 Comparison of the amino acid profiles of fish meal, soybean meal and our Pythium

biomass ...... 42

CHAPTER I: INTRODUCTION

1.1. Omega Fatty Acids

Omega-3 fatty acids are group of long chain polyunsaturated fatty acids (LC PUFAs) which contain a carbon to carbon double at the third position from the terminal methyl group (omega) of the fatty acid chain (Chan, 2009). Omega-3 fatty acids include; Eicosapentaenoic acid (EPA)

C20:5, Docosahexaenoic acid (DHA) C22:6, Docosapentaenoic acid (DPA) C22:5,

Eicosatrienoic acid (ETA) C20:3, and Alpha-linolenic acid (ALA) C18:3 (Rioux Catheline

Bouriel, & Legrand, 1999). The major source of EPA and DHA is fish while ALA can be obtained from nuts and soybeans (Chan, 2009).

Omega-6 fatty acids are a group of long chain polyunsaturated fatty acids (LC PUFAs) which contain a carbon to carbon double bond at the sixth position from the terminal methyl group

(Chan, 2009). Some of the omega-6 fatty acids include; Linoleic acid (LA) C18:2, Gamma- linolenic acid (GLA) C18:3, Eicosadienoic acid C20:2, Dihomo-gamma-linolenic acid (DGLA)

C20:3, Arachidonic acid (ARA) C20:4, Docosadienoic acid etcetera (Rioux et al., 1999). Major dietary sources of omega-6 fatty acids are nuts, vegetable oils, eggs and whole grains.

Diatoms, ubiquitous in marine and fresh waters (Zulu, Zienkiewicz, Vollheyde & Feussner,

2018) are the primary source of omega-3 fatty acids (FAs) for wild-caught fish. Fish and other seafoods are thus considered nutritionally high-quality foods (Strobel, Jahreis &Kuhnt, 2012).

This is due to the presence of high concentrations of omega-3 LC PUFAs in these foods. Fish and other seafoods are thus considered the major sources of these omega fatty acids (Strobel et al., 2012) to human population and farmed fish. The omega fatty acids are essential metabolites especially to human physiological processes such as normal growth and development, prevention 2 of heart or blood vessel diseases, inflammatory diseases, and prevention of cognitive decline by promoting healthy brain functioning (Strobel et al., 2012).

In human beings, omega-3 and omega-6 LC PUFAs are metabolized through the same pathway and therefore they compete for the same enzymes involved in desaturation and elongation of these fatty acids (Strobel et al., 2012). However, each type of the omega fatty acids plays different roles in human physiology and therefore their ratios in the human body influences the overall human health. The preferred ratios are between 1:1 to 5:1 which then becomes highly beneficial health-wise (Strobel et al., 2012). An imbalance to these ratios then leads to terminal illnesses (Strobel et al., 2012).

The US diet entails a high consumption of meats and vegetable oils (Strobel et al., 2012).

These foods are known to have very high levels of omega-6 fatty acids such as linoleic acid which therefore distorts the preferred omega-6: omega-3 ratio. The imbalance leads to a decrease in the levels of omega-3 fatty acids (Cole, Ma, & Frautschy, 2009). Currently dietary ratios of omega-6: omega-3 LC PUFAs in the US diet, stand between 8:1 and 17:1 (Strobel et al., 2012).

This ratio may impair normal brain development hence the recommendations to increase intake of fish providing the necessary omega-3 FAs such as EPA and DHA. The recommended amounts of EPA and DHA is between 200-500 mg/day and thus nutritionists’ advice consuming fish at least twice a week (USDA, 2020). This means that the demand for fish consumption is high.

Fish, just like human beings, are unable to make these fatty acids on their own but depend on the diet they are fed on to provide these essential fatty acids (Strobel et al., 2012) due to lack the enzymes (desaturases) required to synthesize omega-3 PUFAs de novo from an oleic precursor and therefore a dietary intake of linoleic acid and α-linolenic acid is required. In wild-caught 3 fish, algae and diatoms are usually the primary source of omega fatty acids (Strobel et al., 2012).

However, overfishing and pollution of the lakes and seas has led to a decrease in populations of many species of marine fish. With increasing population in the world and the growing demand for consumption of fish from dietary recommendations, fish farming has become a critical component of global food security (FAO, 2020)

Commercial fish farming has risen a thousand-fold since the 19th century while traditional fishing (wild-caught fish) from lakes and oceans has dropped drastically due to overfishing

(Pelletier et al., 2018). Aquaculture has now overtaken the wild-caught fish as a source of proteins and omega fatty acids. 40 essential nutrients comprising of vitamins, amino acids, minerals and fish oil account for the nutritional needs of farmed fish (NOAA_USDA, 2018).

Current fish farms feed their fish on a wide variety of various plant-based ingredients such as legume seeds and oilseed cakes, (Strobel et al., 2012).

The major source of fish oil fed to fish remains to be fish meal (NOAA_USDA, 2018). With the growing trend in aquaculture yet diminishing trend for wild-caught fish, a need for an alternative source of fish oil for fish farming becomes inevitable. Two terrestrially based sources of omega-3 fatty acids are now being introduced to the market. They include omega-3 fatty acids produced by marine algae (Veramaris, 2020) and omega-3 fatty acids produced from genetically modified canola (Ruyter et al., 2019). Both products have high production costs and thus additional sources of omega-3 FAs could potentially be competitive in the marketplace.

1.2. Oomycetes

Oomycetes are a group of filamentous and multicellular eukaryotic organisms which are soil- borne (Kamoun, 2003) pathogens that belong to the kingdom Stramenopila and class Oomycota

(Schroeder et al., 2013). Oomycetes and diatoms are both classified as Stramenopiles, and these organisms have retained the ability to synthesize omega-3 and omega-6 FAs (Zulu et al., 2018).

While they share a hyphal growth habit, the presence of cellulose and β-glycans in their cell wall, coenocytic hyphae and biflagellate zoospores (tinsel and whiplash flagella) are characteristic features that distinguish oomycetes from true fungi (Schroeder et al., 2013).

Species of oomycetes have been identified as pathogens of plants, insects, fish, crustaceans, and microbes. (Kamoun, 2003). Pythium insidiosum is presently the only known pathogen of vertebrate animals. The two most studied genera of oomycetes are Phytophthora and Pythium.

Phytophthora and Pythium species are phenotypically distinct from each other; Pythium exhibit rapid growth on V8 media relative to Phytophthora species which are slow (Wagner et al., 2018). Pythium species are cold loving oomycetes and tend to thrive under low temperature seasons usually below 200C (Wagner et al., 2018). The metabolic requirements between

Phytophthora and Pythium are quite diverse. For example, Phytophthora infestans is hemi- biotrophic while Pythium ultimum which is a good EPA producer (Stredansky, Conti & Salaris,

1999) is necrotrophic (Ah-Fong et al., 2019). Furthermore, Ah-Fong et al (2019) demonstrate that the strategies for using nitrates in P. ultimum are different from that of P. infestans. In P. ultimum, the nitrogen assimilation pathway from nitrate is not feed-back inhibited by amino acid repression as a nitrogen metabolite repressor appears not to be present. Thus, protein synthesis in

Pythium species can be largely supported from inorganic nitrogen sources. 5

1.3. Potential of Pythium to Synthesize Omega Fatty Acids Using Various Sources of

Carbon

Several studies have demonstrated the potential to synthesize the important omega fatty acids in different strains of Pythium (Cantrell & Walker, 2009; Liang et al., 2012; Ren et al., 2017; Lio

& Wang, 2012; Liu et al., 2015; Stredansky, Conti & Salaris 1999). However, Pythium irregulare may be the best producer of omega fatty acids, especially EPA (Liang et al., 2012).

Nevertheless, other strains of Pythium such as Pythium splendens (Ren et al., 2017), Pythium ultimum (Stredansky, Conti & Salaris 1999) and the fungi Umbelopsi ramanniana (Cantrell &

Walker, 2009; Lardizabal et al., 2008) can also be used to synthesize omega-3 fatty acids.

Pythium irregulare produces significant amounts of omega-3 fatty acids, specifically EPA through fermentation with submerged cultures (Cantrell & Walker, 2009). Supplementation of the submerged cultures with 1% glucose {the optimum concentration for maximum biomass production according to (Stinson, Kwoczak, & Kurantz, 1991)} as a source of carbon enhances production of high levels of EPA. Similarly, fermentation of Pythium irregulare by supplying soybean processing co-products as a source of carbon produces ARA and EPA (Lio & Wang,

2012). Soybean cotyledon and soy skim have been employed as potential substrates for Pythium irregulare under different fermentation conditions (submerged and solid-state fermentation) to produce ARA and EPA. Soybean cotyledon was used as a source of carbon in solid-state fermentation and soy skim was used as a source of carbon in submerged fermentation (Lio &

Wang, 2012).

Thin stillage, a by-product of dry milling in corn-ethanol processing plants has been employed as a source of carbon for Pythium irregulare to produce EPA (Liang, Zhao, Strait, &

Wen, 2012). Thin stillage is nutrient rich and can be a carbon, nitrogen, protein source for P. 6 irregulare in the synthesis of EPA (Liang et al., 2012). Cotton seed meal has also been employed as a source of carbon for Pythium irregulare in the production of EPA (Liu, Cao, Guan, Liao &

Cao, 2015). The cotton seed meal supports overall better growth for P. irregulare and with sodium carbonate supplementation, able to produce 245.3mg/L of EPA after 6 days of fermentation at room temperature (Liu et al., 2015).

Pythium splendens strain RBB-5 has also been used to produce EPA as well (Ren, Zhou, Zhu,

Zhang & Yu, 2017). This study elucidated the pathway in P. splendens that is involved in EPA synthesis and the enzymes (desaturases) involved in the biosynthetic process. By application of metabolically engineered strains of P. splendens through amplifying the expression of the enzymes involved in the biosynthetic pathway, introducing an external fatty acid precursor and boosting the fermentation conditions, the amount of EPA produced by these organisms increased significantly (Ren et al., 2017).

Pythium ultimum is also a good producer of LC PUFAs. Stredansky, Conti & Salaris (1999) used P. ultimum in both solid state cultivation and liquid cultivation to produce ARA and EPA.

In this study, cereal grains, malt grains and apple pomace were employed in solid state cultivation while supplementation from 2% glucose was used as a source of nutrition for the oomycete in liquid cultivation. After 6-7 days of growth at 210C, there was a 2.1 mg/g of ARA and 1.14mg/g EPA in solid state cultivation and 0.31mg/g ARA and 0.21mg/g of EPA in liquid cultivation (Stredansky, Conti, & Salaris, 1999). However, in all these studies none has tested the viability of soybean seed-coats as a potential carbon source for Pythium in the production of important omega fatty acids.

1.3.1. Effects of cultural conditions in the biosynthesis of omega fatty acids

Cultural conditions such as growth temperature (Cantrell & Walker, 2009; Liang et al., 2012), moisture content, aeration of samples, incubation time, source and amount of carbon affect the biomass production and the amount and variety of omega fatty acids synthesized (Lio & Wang,

2012). Cantrell and Walker (2009) investigated the effect of temperature in biomass and fatty acid synthesis by P. irregulare through running various experiments at 140C, 210C and 280C.

This study reported maximum fatty acid production at 210C with maximum EPA synthesis at

140C (Cantrell & Walker, 2009) which is consistent with other studies (Jiang & Cheng, 2000;

Wen & Chen, 2001; Wen & Cheng, 2003).

Stinson, Kwoczak, & Kurantz (1991) also investigated optimal conditions for fermentation of

Pythium irregulare for maximum production of EPA. According to this study the maximum biomass production was achieved at 250C while maximal EPA synthesis was achieved at 120C which is consistent with the study by Cantrell and Walker (2009). All these studies suggest that cooler temperatures favor maximum production of omega fatty acids while room temperature is best for maximum biomass production. An addition of glucose boosted the growth rate in this study as well.

The concentration of the nutrient source (carbon source) dictates the amount of fatty acids produced by Pythium species. Liang et al (2012) investigated the effect of thin stillage concentration on EPA synthesis. Increase in the concentration of the nutrient source (beyond

30%) increases the total biomass produced by the Pythium cultures, however, the total EPA yield reduces (Liang et al., 2012). 8

Moisture content influences the optimization of production of maximum omega fatty acids.

Lio and Wang (2012) investigated the effect of moisture content, incubation time, glucose addition, and supplementation with vegetable oil in production of long chain polyunsaturated fatty acids by solid state fermentation (SSF). They noted that limited moisture content in SSF limits the amount of LC PUFAs synthesized. Increase in incubation time relatively increases the amount of EPA and ARA produced (Lio & Wang 2012). Submerged cultures and liquid fermentation perform better overall.

Supplementation of cultures with glucose boosts exponential growth of cultures and in turn synthesis of more LC PUFAs (Lio & Wang, 2012). Another gap in information is that all these studies investigated omega fatty acid production by Pythium using fermentation in liquid and solid cultures. There is no study that has tested the viability of growing Pythium under microaerobic and aerobic cultural conditions in the production of omega fatty acids. This is where our study comes in, to bridge the gap and increase the body of knowledge.

1.4. Research Question

The overarching question throughout this study was; can pathogenic Pythium species isolated from the environment be utilized for bioprocessing of soybean seed-coats to produce a protein and fatty acid source for fish?

1.5. Hypotheses

1. Soybean seed-coats are a viable carbon source for Pythium in the production of omega

fatty acids and proteins

2. High amounts of omega fatty acids production are achieved at cooler temperatures of

around 14-160C 9

3. Aeration of samples and supplementation with 4% sucrose increases omega-3 fatty acid

production

1.6. Goals of the Study

1. To develop a derivatization procedure and a HPLC separation method for major fatty

acids

2. To make and run different fatty acid standards to obtain standard curves to be used

for quantification of fatty acid peaks

3. To optimize the Pythium isolates for maximum fatty acid production

4. To analyze whether sucrose addition and aerobic growth conditions can enhance

growth of Pythium and increase fatty acid levels

5. To quantify the protein levels in the resultant biomass and analyze for their potency

as a protein supplement for aquaculture

CHAPTER II: HPLC ANALYSIS OF FATTY ACIDS

2.1. Introduction

Several methods have been employed in the past for the analysis and quantification of fatty acids after extraction and derivatization. Gas chromatography (GC) being the most popular among them, separates the fatty acids after their conversion to methyl esters (Lima & Abdalla,

2002). However, there are several limitations to this technique. The downside of conversion of the fatty acids to methyl esters step in GC is that methyl esters are also volatile and maybe lost on refluxing during the esterification process (Rioux et al., 1999; Lima & Abdalla, 2002).

Secondly, in GC there is susceptibility of the fatty acids to thermal degradation which in turn leads to structural modification of especially the unsaturated fatty acids (Rioux et al., 1999; Lima

& Abdalla, 2002). The detection method in GC, flame-ionization detection also destroys the fatty acids and thus limits the potential of recovering them for further analysis (Rioux et al., 1999).

High performance liquid chromatography (HPLC) has advantages over GC since the resolved fatty acids are not destroyed but can be collected in fractions to enable additional analysis and is best suited for biological samples (Lima & Abdalla, 2002; Czauderna et al., 2002; Rioux,

Catheline, Bouriel & Legrand, 1999). Another conferred advantage is that the lower temperatures of liquid chromatography protect the unsaturated fatty acids from isomerizing

(Czauderna et al., 2002). HPLC process is also fast, has high resolution, is sensitive to minute amounts of fatty acids and is very specific (Lima & Abdalla, 2002; Czauderna et al., 2002).

2.1.1. Overview of the HPLC process

HPLC is a highly versatile technique that is used to separate the components in a liquid mixture based on their differential interactions with the stationary phase (Petrova & Sauer, 11

2017). It can be used to separate proteins, fatty acids, nucleic acids, polyamines, and other chemical components found in drugs and foods. There are fundamentally two forms of HPLC; the analytical and preparative HPLC. In analytical HPLC, the aim is to identify components making up a liquid mixture by injecting a small volume which is then discarded as waste while in preparative HPLC, the aim is to purify a mixture and the desired amounts of each component are collected in fractions, which are then concentrated by removal of solvents.

HPLC is a modification of column chromatography. In classical column chromatography, the stationary phase is a column packed with microscale beads. These beads are functionalized with chemical groups that promote the interaction between the stationary phase (beads in the column) and mobile phase (components of the liquid mixture) (Petrova & Sauer, 2017). The interaction differs between the different components making up the mixture in the mobile phase and hence their elution times from the column of beads varies. Once the component travels through the column it passes through a detector which records the amount of time taken by the component to be eluted from the column and reach the detector (Petrova & Sauer, 2017). This time is known as the retention time. The detector is then connected to a computer software that plots a graph

(chromatogram) of retention time versus intensity.

The retention time is specific for different components and therefore its used to identify a component in the liquid mixture in relation to an already known pure standard while the peak size or area under the peak is used to quantify the amount of substance in the liquid mixture. The choice of a stationary phase is very crucial, and it depends on the hydrophobicity or hydrophilicity of the liquid mixture components (Petrova & Sauer, 2017). In the past cellulose powder was often used, and sometimes alumina is common. The most popular columns today are 12 constructed with alkyl chains bounded to silica particles so that the stationary phase is strongly hydrophobic.

2.1.2. Derivatization of fatty acids as Naphthacyl derivatives

Fatty acids are weakly chromophoric and therefore their detection via UV is not very sensitive

(Lima & Abdalla, 2002). To boost the detectability of fatty acids, several chromophoric groups have been used as derivatizing agents. These include; methyl esters (Carvalho et al., 2012), nitro- benzyl, benzyl, naphtha-iminoethyl esters, phenacyl esters (Umeh, 1971; Chen & Anderson,

1992), phenyl-azo-phenacyl (Umeh, 1971), and Naphthacyl esters (Rioux et al., 1999; Cooper &

Anders, 1974). The chromophore 2-bromoacetophenone (DBAP) is the most popular chromophore since it amplifies the sensitivity of detectability via UV allowing up to nanogram amounts of fatty acids to be detected (Rioux et al., 1999).

2.2. Materials and Methods

2.2.1. Raw materials

Soybean seed-coats are the raw materials that were used in this project as a carbon source for

Pythium. The soybean seed coats were obtained from the soybean council from the cargo soybean processing plant in Sydney, Ohio. Approximately 6 million metric tons of soybean seed coats are produced annually (O’Brian et al., 2014). They are low cost, low value soybean by- products from soybean processing that are usually fed to cattle as a fiber supplement. They are an underutilized resource that has potential benefits and one of our main aims is to turn these valueless and less nutritious soybean seed coats into a product of value in the production of essential omega-3 and omega-6 fatty acids.

2.2.2. Reagents and chemicals

The chemicals used in this study were purchased from various sources. Methanol (General use HPLC-UV grade) and ethanol (190 proof) were obtained from PHARMCO-Greenfield

Global (Brookfield, USA). Acetonitrile-190 was bought from CALEDON laboratory chemicals limited (Georgetown, ON, Canada). Liquid chloroform environmental grade 99.8% was purchased from Alfa Aesar (Ward Hill, MA, US). Hexane, Acetone (UN1090), potassium hydroxide (KOH), and Triethylamine (TEA)- HPLC Grade were purchased from Fisher

Scientific (Fair Lawn, NJ, USA). Hydrochloric acid (HCL) and Acetic acid were purchased from

EMD Millipore corporation/EMD chemicals Inc (Billerica, MA, USA). Gamborg B5 medium

(basal salt mixture) was purchased from Duchefa Biochemists (Haarlem, Netherlands). V8 juice was bought at a local store. The fatty acid standards; Eicosadienoic acid, EPA, DHA, DPA, myristic acid, Dihomo_γ_linolenic acid, 3_hydroxy_octanoic acid, and docosanoic acid were purchased from CAYMAN CHEMICAL COMPANY (Ann Arbor, MI, USA). DBAN, ARA and

γ_linolenic acid were purchased from SIGMA ALDRICH (St. Louis, MO, USA). Lauric acid, palmitic acid, stearic acid, caprylic acid, and behenic acid were purchased from Nutritional

Biochemicals Corporation (Cleveland, OH, USA).

2.2.3. Isolation and propagation of Pythium

Soil samples were collected from low-lying areas of agricultural fields around Bowling

Green, Wood County, Ohio. Soybean seeds were then germinated in the wet soil and as they germinated, they were infected by Pythium present in the soil. The infected tissue was sliced out and placed on selective V8 media amended with Benomyl 100mg/L, pentachloronitrobenzene

100mg/L, pimaricin (0.2% [wt./vol]), rifampin (10mg/L), and ampicillin (100mg/L). Pythium isolates were readily distinguished from Phytophthora species by their rapid growth on V8 14 media. The Pythium isolates used were labelled WC001, WC002, WC003, WC004, WC005,

WC006, and WC007 respectively.

The cultures of different Pythium isolates from the seven different areas were then maintained on V8 media and periodically transferred to new plates every three weeks. To eliminate contamination from bacteria, the V8 propagation media was augmented with antibiotics;

Valinomycin (1μl/ml of V8 media), Rifampicin (1μl/ml of V8 media), Tetramycin (1μl/ml of V8 media), and Ampicillin (1μl/ml of V8 media) until the cultures were visibly free of microbes.

The protocol used for making V8 media is as follows: For a volume of 1L, V8 media was prepared by adding 200ml V8 juice, 3g of calcium carbonate, and 15g of agar to 800ml of distilled water and mixed well. The contents were then autoclaved at 1210C for 15 minutes, let to cool down to around 500C, added antibiotics (1μl/ml of V8 media) and poured on plates. To transfer the culture, a small plug of actively growing Pythium mycelia was cut out of the initial plate using a sterile scapel and introduced onto a new Petri dish containing fresh V8 media. The fresh cultures in petri-dishes were incubated upside down at room temperature for three days, and then stored at 160C for a maximum of three weeks before transferring again.

2.2.4. Cultivation of Pythium isolates

The Pythium plugs from different isolates on V8 plates and soybean seed-coats were first cultured under hypoxic conditions in liquid Gamborg B5 (GB5) media. To prepare the cultures;

7 clean 1L flasks with rubber corks were assembled, 5g of soybean seed coats were weighed into each of the 7 clean flasks. Gamborg B5 media was prepared by weighing out 2.147g of GB5 salt and dissolving it in 700ml of diH20. Then 100ml of GB5 solution was aliquoted into each of the

7 flasks and the mixture autoclaved at 1210C for 15 minutes. 15

After cooling the flasks with GB5 media and soybean seed coats to room temperature, the inoculum was prepared by transferring 5 Pythium mycelial plugs of approximately 0.2cm2 each from V8 plates into the respective 1L flasks. Half a milliliter of vitamins added into each flask to boost growth and the flasks sealed using silicone sponge closures ideal for hypoxic conditions.

The stock concentration of the vitamins used was made according to the protocol by (Fang et al.,

2017); 10μl of 0.02 g/ml Biotin, 10μl of 0.02 g/ml Folic acid, 0.012g of L_inositol 0.06g

Nicotinic acid, 0.18g Pyridoxine-HCL, 0.015g Riboflavin, 0.38g Thiamine-HCL, water up to the

300ml mark, filter sterilize and store at 40C.

The Pythium cultures were grown under varying temperature conditions starting with room temperature in a shaker from 2 to 5days and then transferred to lower temperatures from 24 hours to 3 days (see Table 2). Growth conditions were varied from longer growth periods at room temperature and shorter periods at colder temperatures with the aim of determining the optimal temperature conditions for maximum production of biomass and fatty acids.

2.2.5 Optimization of growth and FA production by Pythium

To optimize for the best temperature combinations for faster growth of the Pythium

isolates and maximum fatty acid production, we did different trial runs; the first trial

involved growing the isolates for 2 days in the shaker at room temperature (RT) and 4

days in the cold room (CR). In the second trial, isolates were grown for 4 days at RT and 2

days in the CR. In the third trial, isolates were grown for 6 days at RT and 3 days in the

CR. In the fourth trial; 5 days at RT and 3 and half days in the CR. In the fifth trial;

isolates were given 2 days at RT and 2 days in the CR, and in the sixth trial; 5 days at RT

and 3 days in the CR. The biomass was then harvested after each set of growth conditions,

frozen overnight at -200C, dried up for three days at 650C and pulverized for downstream 16

processing in fatty acid extraction and quantification using High Performance Liquid

Chromatography (HPLC).

2.2.6. Fatty acid extraction

The Acidic Bligh and Dryer fatty acid extraction protocol was employed in the extraction and release of fatty acids for HPLC analysis (Jensen, 2008). The harvested and dried Pythium biomass was pulverized by a coffee grinder to ensure uniform access of small particles to organic solvents. The biomass of each isolate weighing approximately 100mg was then transferred into

8ml vials, 1 ml of 3M HCl added, and the samples heated for 1 hour at 80°C in a dry hot bath.

Next 1.50 ml methanol and 1.00 ml chloroform plus the internal standard-undecanoic acid (10 μl of 500μM undecanoic standard) were added and this mixture was shaken for 1 min. Then 1 ml of deionized water and 2.00 ml chloroform were added, the tube shaken for 1 min, and samples allowed to settle. 1.5 ml of lower chloroform layer was removed with an Eppendorf pipette and transferred into new clean 8ml vials and the samples were dried down in a nitrogen airstream.

2.2.7. Saponification

The purpose of this step is to break down the bonds connecting fatty acids to glycerol. An aliquot (1.5-2 ml) of the extracted fatty acids that was dried under nitrogen, was immediately re- suspended in 2 ml of 2.0% potassium hydroxide (KOH) in ethanol, and saponified at 100°C for

30 minutes in a dry bath. Precaution was taken to ensure the dry bath was up to the required temperature before adding the KOH. After cooling to room temperature, 1 ml of deionized autoclaved water and 100µl of concentrated hydrochloric acid (HCL) were added and mixed well. The released free fatty acids were then extracted twice with 2 ml of hexane and dried under a nitrogen stream. The released fatty acids were then derivatized as Naphthacyl derivatives as described below. 17

2.2.8. Goal 1: A derivatization procedure and HPLC separation method

In this study, we achieved our first goal by using Naphthacyl derivatives as our derivatization technique, a method developed by Rioux et al (1999) and modified in our lab. In this method, the chromophore 2-bromo2'-acetonaphthone (DBAN), which boosts sensitivity even more at 246nm

UV detection, was used to derivatize fatty acids after extraction and saponification. The released and dried fatty acids from the saponification step were then taken up in 2ml of ethanol and a

200μl aliquot of this stock solution of fatty acids was then used at the derivatization step.

To convert the liberated fatty acids to Naphthacyl esters, 750μl of DBAN (1.2g/25ml solution of acetone) and 100μl of Triethylamine (TEA) were added to the 200μl aliquot of the fatty acid extract at the derivatization step. The mixture in 8ml screw top vials was then incubated in a dry bath for 40 minutes at 650C. The reaction was stopped by adding 100μl acetic acid in acetone (2 grams A.A./L acetone) and dried down under a stream of nitrogen gas. The dried fatty acid

Naphthacyl esters were then re-suspended in 1 ml of acetonitrile or methanol/dichloromethane

3:1, filtered using 0.45-micron syringe filter, and transferred to sample vials for HPLC analysis.

2.2.9. Separation of fatty acid Naphthacyl esters by HPLC

We employed reverse-phased analytical high-performance liquid chromatography to identify the different types of fatty acids produced by different Pythium isolates. Injection volumes of 15-

50μl of the derivatized fatty acid Naphthacyl esters were separated on Agilent 1120 Compact

HPLC using Gemini 3-micron NX-C18 110Å 250mm C-18 column. These small volumes were used to enable quantification of small peaks while ensuring the peak absorbance of larger peaks did not exceed 4000mAU beyond which the peak area approximation goes out of scale. The small pore size of this column enables a high level of resolution but typically operates at >220 18 bars. The column temperature was maintained at 300C. The elution of fatty acids was programmed at a flow rate of 1ml/min with a gradient of solvent A(10%ACN/90% water)/solvent B(85%methanol/15%ACN) starting at 20:80 (v/v) increasing linearly to 9:91 (v/v) in 30 minutes, then further increasing linearly to 0:100 (v/v) in 10 minutes, holding at 0:100 (v/v) for 5 minutes and then returning to initial conditions in 5 minutes . The elution was tracked by a

UV detector at 246nm, which is the maximum absorbance for Naphthacyl esters. A chromatogram was then generated by the Agilent Lab Advisor Software- B.02.14 connected to the HPLC system. Each sample was resolved for 60 minutes. Resolution was achieved by the chromatograms generated by different retention times. 19

CHAPTER III: RESULTS AND DISCUSSION

3.1. Goal 2: Fatty Acid Standard Curves

Fatty acid standards are highly purified salts of individual fatty acids with already known concentrations and replicable retention times during HPLC analysis. The retention times (in minutes) of the analyzed fatty acid standards are as follows; 11.5 (caprylic acid), 21.8

(undecanoic acid), 32.6 (EPA), 33.9 (alpha linolenic acid), 34.2 (myristic acid), 35.8 (DHA),

36.6 (ARA), 37.9 (linoleic acid), 38.2 (DPA), 39.1 (Eicosatrienoic acid), 39.2

(Dihomo_γ_linolenic acid), 39.9 (palmitic acid), 40.9 (oleic acid), 41.7 (Eicosadienoic acid), and

43.5 (stearic acid). These retention times have a 0.1%-0.4% standard deviations depending on the accuracy of the measurement ratios of polar and non-polar solvents.

Figure 1 below shows derivatization and HPLC analysis of fatty acid standards of different volumes. 10μl, 20μl, 30μl, 40μl, and 50μl of each fatty acid standard was derivatized (taking note of each fatty acid stock concentration) and eluted through HPLC. The corresponding areas for each volume were plotted as areas under peaks versus the weight of fatty acids at injection

(15μl) for each derivatized volume. Simple linear regression analysis was done using Prism 8.0 software (GraphPad software San Diego CA).

Eicosapentaenoic acid Eicosadienoic acid

5×107 1.5×108

4×107 1×108

7 s

3×10 s

r A 2×107 A 5×107 1×107 A B

0 0 0.0 0.1 0.2 0.3 0.4 0.5 0.0 0.5 1.0 1.5 Mass in g present in 15l Mass in g present in 15l

Undecanoic acid Dihomo__linolenic acid

1.5×109 6×107

1×109 4×107

A A 5×108 2×107 C D

0 0 0 20 40 60 0 10 20 30 40 Mass in g present in 15l Mass in g present in 15l

Docosahexaenoic acid Myristic acid

1.5×108 4×108

3×108

1×108

a e e 8

r 2×10

A A 5×107 1×108 E F

0 0 0.0 0.5 1.0 1.5 0.0 0.5 1.0 1.5 2.0 Mass in g present in 15l Mass in g present in 15l

Arachidonic acid Docosapentaenoic acid 1×109 4×108 8×108 3×108 6×108 2×108

Areas 8

4×10 Areas

8 1×108 2×10 G H

0 0 0 20 40 60 0 2 4 6 8 Mass in g present in 15l Mass in g present in 15l

Stearic acid Palmitic acid

3×108 3×108

2×108 2×108 Areas Areas 1×108 1×108 I J

0 0 0.0 0.5 1.0 1.5 2.0 0.0 0.5 1.0 1.5 2.0 Mass in g present in 15l Mass in g present in 15l

Figure 1. Fatty acid standard curves. A) Standard curve for Eicosapentaenoic acid (EPA) B)

Standard curve for Eicosadienoic acid C) Standard curve for Undecanoic acid D) Standard curve for Dihomo_γ_linolenic acid (DGLA) E) Standard curve for Docosahexaenoic acid (DHA) F)

Standard curve for myristic acid G) Standard curve for Arachidonic acid (ARA) H) Standard curve for Docosapentaenoic acid (DPA) I) Standard curve for stearic acid J) Standard curve for palmitic acid. Each individual fatty acid standard had a unique concentration depending on the stock solution and on the amount required to project a straight line. 22

Figure 2 below shows a chromatogram of a standard fatty acid mixture. 11 peaks were identified based on the retention time of the available fatty acid standards. Our results are consistent with those of Rioux et al (1999) for elution through the reverse-phased column and therefore retention time increases with long chain fatty acids. The order of elution can hence be predicted using FA carbon chain length alone, however, unsaturated fatty acids are eluted earlier than saturated fatty acids, for example, EPA (C20:5), a long chain polyunsaturated fatty acid is eluted at 32.6 minutes while stearic acid (C18), a saturated fatty acid is eluted at 43.5 minutes.

Some of the retention times of fatty acids that are closely resolved overlap with each other and therefore are coeluted, for example, Eicosatrienoic acid (an omega 3 FA) resolved at 39.1 minutes and Dihomo_γ_linolenic acid (an omega 6 FA) at 39.2 minutes overlap. Also, ALA resolved at 33.9 minutes and myristic acid at 34.2 minutes sometimes overlap and therefore necessitates confirmation with LCMS to fully resolve the identity of these FAs peaks. In analyzing a complex mixture of fatty acid standards, Rioux et al (1999) was not able to resolve positional isomers of octadecenoic acid using this method. 23

Figure 2. Chromatogram of a mixture of fatty acid standards. Peaks: 11.5 (caprylic acid), 21.8

(undecanoic acid), 25.4 (lauric acid), 32.6 (EPA), 34.2 (myristic acid), 36.6 (ARA), 37.9

(linoleic acid), 38.2 (DPA), 39.2 (Dihomo gamma linolenic acid), 39.9 (palmitic acid), 40.9

(oleic acid), 41.7 (Eicosadienoic acid), and 43.5 (stearic acid). 24

Table 1. Resolution of derivatized fatty acids using Gemini 3-micron NX-C18 110Å 250mm column Retention time Common name Lipid Chemical name-IUPAC (RT) number 11.5 Caprylic acid 8:0 Octanoic acid 21.8 Undecanoic acid 11:0 Undecylic acid 25.6 Lauric acid 12:0 Dodecanoic acid 32.6 Eicosapentaenoic acid 20:5 all-cis-5,8,11,14,17-eicosapentaenoic acid 33.9 α_linolenic acid 18:3 all-cis-9,12,15-octadecatrienoic acid 34.2 Myristic acid 14:0 1-tetradecanoic acid 35.8 Docosahexaenoic acid 22:6 all-cis-4,7,10,13,16,19-docosahexaenoic acid 36.6 Arachidonic acid 20:4 5,8,11, 14-Eicosatetraenoic acid 37.9 Linoleic acid 18:2 Cis- 9, 12-octadecadienoic acid 38.2 Docosapentaenoic acid 22:5 all-cis-7,10,13,16,19-docosapentaenoic acid 39.1 Eicosatrienoic acid 20:3 Icosa-8, 11, 14- trienoic acid 39.2 Dihomo_γ_linolenic acid 20:3 8, 11, 14-Eicosatrienoic acid 39.9 Palmitic acid 16:0 Hexadecanoic acid 40.9 Oleic acid 18:1 Cis-9-octadecenoic acid 41.7 Eicosadienoic acid 20:2 11, 14-dienoic acid 43.5 Stearic acid 18:0 Octadecanoic acid 44.1 Unknown FA 46.5 Unknown FA The rows highlighted in blue are omega-3 FAs and the rows highlighted in green are omega-6 FAs. Unresolved fatty acids such as Eicosatrienoic acid and DGLA require LCMS to confirm the identity of the fatty acids. Unhighlighted rows represent saturated fatty acids.

3.2. Analysis of Fatty Acid Profiles from the Isolates

Next we derivatized and HPLC analyzed samples from our environmental Pythium isolates.

We used Naphthacyl esters and reverse phased analytical HPLC method to determine the major fatty acid peaks produced by the Pythium isolates given soybean seed coat hulls as a Carbon source (figure 3.) Four omega-3 fatty acids (EPA, ALA, DPA and ETA) and three omega-6 fatty acids (DGLA, ARA, EDA) were detected. DGLA and ETA co-eluted as an unresolved major peak. Other major peaks were those of saturated fatty acids; palmitic acid, stearic acid, and oleic acid. The fatty acid profile from our isolates generates late peaks at around retention times of 44.

1, 46.5, 48.0 and 50.0 minutes that we have yet to identify, but we hypothesize that peaks eluted after stearic acid C18: 0 could be long chain saturated fatty acids; arachidic acid (C20:0) and 25 behenic acid (C22:0) since unsaturated fatty acids tend to be eluted earlier. Below are figures 3 and 4 that show the fatty acid profiles from two of our isolates; WC005 and WC007.

Figure 3. Chromatogram of HPLC elution of isolate 5 (WC005)

Figure 4. Chromatogram of HPLC elution of isolate 7 (WC007). The peaks with an ‘unknown’ are the unidentified peaks, however we assume the late peaks could possibly be long chain saturated fatty acids.

To compare the fatty acid profiles of the biomass from our Pythium isolates with that of fish oil, we derivatized a sample of fish oil pill from a nutrition supplement (mega-red omega-3 supplement pills) of extracted sardine and salmon oil. We employed the same protocol for extraction, derivatization and HPLC analysis of the fish oil pill. Comparison of the two profiles shows that our fatty acid profile is comparable to that of fish oil and has similar complexity and variety of omega-3 fatty acids. 27

Figure 5. Chromatogram overlays of the fatty acid profiles of sardine oil (fish oil) and isolate

WC007. The fatty acid profile resulting from Pythium growth assays with soybean seedcoats as the main carbon source mimics that of fish oil/sardine oil as shown in this figure.

3.3. Goal 3: Optimization of Omega-3 FA Production in Pythium

To optimize for the isolate best suited for maximum production of omega-3 fatty acids, we grew the isolates under different combinations of temperature shifts. Growing the Pythium isolates at room temperature promotes faster biomass production while shifting the temperature to cold growth chambers at 160C promotes omega fatty acid production (Liang et al., 2012).

Under cooler temperatures then cells switch to synthesizing more of unsaturated fatty acids to 28 keep the fluidity of their cell membranes intact since unsaturated fatty acids characteristically remain liquid at lower temperatures than saturated FAs.

Analysis of the samples grown under different growth conditions show that the samples grown under hypoxic conditions for 5 days in the shaker at room temperature and 3 days in the cold room had the highest total biomass and fatty acid levels as shown below in Table 2. Thus, growth for 5 days at room temperature and 3 days in the cold room were determined as the best possible set of growth conditions to yield maximum amounts of omega fatty acids.

Table 2. Temperature shift combinations for optimization of omega-3 fatty acid production Growth No. of Days in No. of 500ml/min Addition Average Total Condition the shaker at Days in O2 of 4% % μg/mg 250C cold room bubbling Sucrose biomass dry at 160c weight of FA 1 2 2 X X 45.7 - 2 2 4 X X 68.5 - 3 4 2 X X 61 - 4 5 3 X X 72 8.48 5 5 3 1/2 X X 76 - 6 6 3 X X 61.7 - 7 4 3 X X 69.4 - 8 3 4 √ X 74.1 - 9 5 3 X √ 74.3 61.56 10 5 3 √ √ 101.9 94.40 The conditions of growth highlighted in blue in the table above have been determined to be the best conditions so far to produce maximum amounts of fatty acids. Column 1 represents the various growth conditions. Column 2 indicated the number of days the isolates were grown in a shaker at room temperature. Column 3 shows temperature shift and the number of days the isolates were left in cold growth chambers. In column 4 and 5; an X means oxygen and sucrose were not added while a √ shows that oxygen and 4% sucrose were added to the cultures correspondingly. Column 6 represents the average percentage biomass for all the isolates grown under a specific set of growth conditions. Column 7 shows the corresponding total μg/mg FA for the optimized growth conditions.

3.4. Goal 4: Effect of Sucrose and Oxygen Supplementation on FA Concentration in the

Pythium Biomass

In our pilot experiments, 5g of soybean seed coats were autoclaved in 100 ml of Gamborg B5 nutrient solution using l liter flasks and inoculated with plugs of Pythium cultures from V8 plates after addition of supplemental vitamins. The flasks were stoppered with porous silicon stoppers.

Due to the porous nature of the silicon lids, it is likely that cultures were growing under hypoxic conditions.

The total fatty acid levels of extractable fatty acids from dried biomass was achieved by weighing out 500mg of dry biomass and extraction of the fatty acids proceeded following the

Acidic Bligh and Dryer fatty acid extraction protocol explained in the methodology section, however, we doubled the volumes of extraction reagents since we had doubled the mass by weight being extracted. The amount of FA was assessed as gravimetric weight relative to dry weight of the Pythium biomass. The percentage by mass of fatty acids was 2.76% for the samples grown under hypoxia. This value is consistent with the values we get using standard curve slope to calculate fatty acid content in the same samples.

To test whether growth and FA production might be enhanced by the addition of a readily digestible carbon source, we supplemented the Gamborg B5 medium with 4% sucrose and bubbled 500ml/min of oxygen into the flasks. Figure 6 below shows the experimental set up for oxygen and sucrose supplementation and the resultant biomass after this growth condition. When we allowed luxury consumption of carbon by the organisms (Pythium) by reducing the carbon stress at the start of the growth assays through adding 4% sucrose and bubbling oxygen through the samples; the weight of extractable fatty acids doubles to 5%. This analysis was done on 30 isolate WC007. Since FA production increased with the addition of sucrose and oxygen supplementation, we concluded that under hypoxic conditions the isolates were carbon limited.

A B

Figure 6. Experimental set up during supplementation of 4% sucrose and bubbling oxygen. A)

Flasks showing resulting biomass from the set up B) Flasks showing the set up for oxygen bubbling into the samples using Compact 525DS Oxygen Concentrator at a rate of

500ml/minute. Oxygen was bubbled through deionized autoclaved water to maintain the moisture content of our samples during the growth period.

Figure 7 below shows that we get bigger peaks for omega-3 fatty acids with addition of 4% sucrose at the start and bubbling of oxygen through the samples during growth assays. There are also extra peaks of omega-3 fatty acids that were not present under hypoxic conditions. Bubbling of oxygen into the samples promotes faster growth rate and formation of more biomass.

Supplementation from 4% sucrose and oxygenation of our isolates during the growth also leads to an increase in an unknown late peak after stearic acid.

Figure 7. Chromatogram of HPLC elution for WC004 and WC007 under 4% sucrose and oxygen supplementation. The peaks with a question mark are unknown peaks yet to be identified. 32

3.5. Quantification of Fatty Acids

To quantify the total amount of fatty acids present in the Pythium biomass from our isolates, we used the gradient values from the standard curves. The standard curves were generated with already known concentrations and used to estimate the total amounts of FAs for the peaks generated from processing our isolates. The total amount of fatty acids increases by 8-fold with the addition of 4% sucrose and bubbling of oxygen through the samples. This suggests that the production of fatty acids using Pythium is optimizable.

Table 3. Analysis of fatty acids in μg/mg dry weight of cultures in flasks under hypoxic conditions for isolate WC007

RETENTION PRODUCT NAME g/mg dry TIME (mins) *- omega-3 weight **-omega-6 Areas/mg 32.7 EPA* 12963977 0.13 33.7 α-linolenic* 16822132 0.17 35.5 DHA* 13590870 0.14 36.7 ARA** 11296069 0.68 37.5 Linoleic** 87529411 5.26 39.8 Palmitic 132768831 0.98 40.7 Oleic 82773335 0.61 41.8 Eicosadienoic** 7420300 0.073 43.6 Stearic 47227458 0.35 Totals 8.48 The * in the table above signifies omega-3 FAs and ** represent omega-6 FAs

Table 4. Analysis of fatty acids in μg/mg dry weight for isolate WC007 under 4% sucrose supplementation

PRODUCT NAME RETENTION *- omega-3 g/mg dry weight TIME (mins) **-omega-6 Areas/mg 32.5 EPA* 132682752 1.36 33.7 α-linolenic* 232411882 2.38 35.4 DHA* 292971352 3.04 36.5 ARA** 84003506 5.05 37.3 Linoleic** 347097162 32.25 38.9 Eicosatrienoic* 41475617 3.85 39.8 palmitic 462658449 5.26 40.6 oleic 510430910 5.79 43.5 stearic 118859383 1.37 Total 61.56 The * in the table above signifies omega-3 FAs and ** represent omega-6 FAs. The tables summarizes only those peaks which we have identified FA standards for.

Table 5. Analysis of fatty acids in μg/mg dry weight isolate WC007 under 4% sucrose and oxygen supplementation

PRODUCT NAME RETENTION *- omega-3 g/mg dry weight TIME (mins) **-omega-6 Areas/mg 32.5 EPA* 246617705 2.53 33.6 alpha linolenic* 549255917 5.63 35.3 DHA* 314700075 3.27 36.5 ARA** 149124110 8.96 37.3 Linoleic** 463742615 27.86

38.9 DGLA/Eicosatrienoic** 57019164 35.33 39.8 palmitic 588521990 4.32 40.6 oleic 620627302 4.56 41.5 Eicosadienoic** 26167823 0.26 43.5 stearic 224013217 1.67 Totals 94.4 The * in the table above signifies omega-3 FAs and ** represent omega-6 FAs. This table summarizes only those peaks that we have confirmed identity for.

The graphs below show a comparison of the relative amounts of individual omega-3 fatty acids in the isolates grown under hypoxic conditions without supplementation with sucrose and relative amounts of indiviual omega-3 fatty acids in the isolates under supplementation with 4% sucrose and 500ml/min oxygen. 34

After supplementation with 4% sucrose the omega-3 fatty acid peaks increased significantly, however in the analysis of individual peaks DPA has been lost. This is an unexpected outcome, further studies are required to figure out the reason why we were able to achieve this observation. Although it could be scientifically explained as DPA is an intermediate between

EPA and DHA (Ritcher & Kris- Etherton, 2016), chances are the DPA that was synthesized was all convereted to the end product DHA in a nutrient rich environment.

Figure 8. Relative amounts of individual omega-3 fatty acids in the isolates under hypoxic conditions

Eicosapentaenoic acid Alpha_linolenic acid Docosapentaenoic acid 8 8 2.5×10 6×10 8×108 WC001 WC001 WC001 8 2×10 WC002 WC002 WC002 6×108 WC003 4×108 WC003

8 WC003 s

s 1.5×10

a a

WC004 WC004 a e

e 8 WC004

e r

r 4×10

r A A 8 1×10 WC005 WC005 A WC005 2×108 WC006 WC006 8 WC006 5×107 2×10 WC007 WC007 WC007 0 0 0 1 2 3 4 5 6 7 1 2 3 4 5 6 7 1 2 3 4 5 6 7 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 C C C C C C C C C C C C C C C C C C C C C W W W W W W W W W W W W W W W W W W W W W Pythium isolate Pythium isolate Pythium isolate

Figure 9. Relative amounts of individual omega-3 fatty acids in the isolates under supplementation with 4% sucrose and 500ml/min oxygen 35

3.6. Stability of the Naphthacyl Derivativess

To analyze for degradation of the Naphthacyl derivatives over time under light, I derivatized the released fatty acids from isolate WC007 and run them on HPLC at an interval of every 12 hours for three days and compared areas of similar sized injection volumes over the time period.

The samples were not kept in a dark place but were left in the autosampler over the experimental period. We assumed at time zero, the areas generated by the chromatogram are 100% of the stable derivatives before degradation. Then we calculated the percentage deviations from time zero up to 72 hours to measure for degradation. Table 4 below shows the percentage deviations.

Table 6. Stability analysis of Naphthacyl derivatives of fatty acids

Time (h) Fatty acid Peak areas 0 12 24 48 72 Undecanoic 608908552 100 108.9 109.2 108.3 107.9

Unknown 64629077 100 128.8 128.8 128.8 130.1 EPA 54962236 100 102.4 102.8 104.7 105.8 Alpha linolenic 126372815 100 103.4 103.1 103.9 105.2 DHA 136758520 100 100.8 101.2 102.3 103.5 ARA 46169415 100 100.1 101.4 102.4 103.6 linoleic 166957301 100 100.3 100.8 101.7 102.6 Dihomo_γ_linolenic 25614431 100 102.1 108.5 110.9 117.2 Palmitic 308326303 100 102.9 100.7 101.4 102.4 Oleic 450177714 100 98.3 98.6 98.2 99.9 Stearic 73031422 100 100 99.9 100.4 101.4

Fatty acids are listed in the elution order of derivatives. Equal volume injections (50μl) were made from the same sample at 0 hours, 12 hours, 24 hours, 48 hours and 72 hours. Peak areas were calculated automatically using Agilent Lab Advisor software. Peak areas at various times following the initial injection are expressed as a percentage area at time zero. The deviations of percentage areas are around 1-2% which is not significant enough to show degradation. This time-dependent degradation experiment of our Naphthacyl derivatives show that the derivatives are stable within a three-day period if left at room temperature and semi-dark conditions. For long-term stability they should be kept at -200C under dark conditions.

3.7. Reproducibility

The repeatability of the derivatization technique modified in our lab was determined by extracting FAs from isolate WC007 biomass and dividing the released fatty acids into aliquots which were maintained at -200C. The aliquots were derivatized independently after a time span of 72 hours. Each aliquot derivatized was run through HPLC and their areas compared. The areas were found to be consistent among the various aliquot replicates.

Table 7. Reproducibility of the derivatization technique

Fatty acid Peak areas Peak areas Ratio of areas 1st derivatization 2nd derivatization

Undecanoic 642147151 619979348 1.04

EPA 57783584 56968647 1.01

Alpha linolenic 132629347 130496988 1.01

DHA 142061683 138522032 1.03

ARA 47830198 46735121 1.02

Linoleic 173634539 167916671 1.03

Dihomo gamma linolenic 26980784 24771958 1.09

Palmitic 319269808 307599487 1.03 oleic 455563199 435588178 1.05 stearic 70595510 79388415 0.89

3.8. Goal 5: Amino Acid Analysis

Proteins are an important component of fish feed (Jobling, 2016) and thus the quantification and composition analysis of the amino acid profile in our Pythium biomass was paramount.

Classical protein source for aquaculture has been fish meal and animal residues (Jobling, 2016) 37 but due to diminishing wild-caught fish, plant-based sources of proteins are gaining popularity.

Although plant-based protein sources are cheap to acquire, they are lacking in some of the essential amino acids, taurine and cholesterol which are vital for normal physiological functioning of fish and may also contain other anti-nutritional factors which are indigestible and thus undesirable (Jobling, 2016). Plants as protein sources for aquaculture suffer from a deficiency of methionine and lysine (Nunes et al., 2014; Jobling, 2016). However, with improved technologies and supplementation of deficient amino acids, plant-based protein sources are a feasible fish feed alternative.

To estimate the total amino acid content in our Pythium biomass; 100mg of biomass from five samples (WC001, WC003, WC004, WC005, and WC007) grown under hypoxic conditions without sucrose supplementation were sent to Creative Proteomics (45-1 Ramsey Road, Shirley,

NY 11967, USA) for acid hydrolysis and amino acid quantification. Aliquots of the samples were hydrolyzed using 0.5% phenol in 6N HCL. The final dried pellets from the samples were then dissolved in 20mM HCL and derivatized. The amino acid levels were quantified using

Agilent 1290 Infinity II UPLC. The concentrations in pmol/μl and in nmoles/mg for each amino acid detected in the samples are shown in the table attached in appendix 1. Note: Cysteine and

Methionine are oxidized into Cya and MetSO2 respectively. Glutamine and Asparagine are converted to Glutamic acid and Aspartic acid, respectively. Tryptophan is destroyed during hydrolysis.

Table 8 below shows a summary of the individual amino acids in mg/g dry weight for each of the isolates analyzed. Total amino acid levels in the samples ranged from 20.35 to 25.9% (Table

8). The metSO2 content in our Pythium biomass range from 0.2-0.39% while lysine is in the ranges of 1.9-2.1%. Fish and shrimp require 0.5- 1.5% and 0.7-0.9% of dietary methionine 38 respectively. The methionine content in our biomass require supplementation to be on the required levels in fish feed. Fish require 1.2-3.3% of lysine in their diet while shrimp requires

1.6-2.1% (Nunes et al., 2014). Our lysine content is within the required levels. The amino acid profile of the Pythium biomass shows that the relative levels of most amino acids is high enough that it serves as a potential protein source for fish feed that does not require further supplementation of these essential amino acids. Although different fish species have different levels of amino acid requirements, our AA profile seems adequate for most fish species.

Variation in protein levels could be due to growth conditions, genetic variation of samples or contamination of oomycete biomass with undegraded soybean hulls.

Table 8. Summary of the amino acid profile in the resulting Pythium biomass

WC001 WC003 WC004 WC005 WC007 mg/g g/100 mg/g g/100g mg/g g/100g mg/g g/100g mg/g g/100g AA dry wt. g N dry wt. N dry wt. N dry wt. N dry wt. N Cya 4.04 2.0 4.39 1.7 4.51 2.0 4.038 2.0 4.45 2.0 His 7.61 3.7 9.13 3.5 8.29 3.6 7.69 3.8 8.29 3.7 Ser 14.51 7.1 17.33 6.7 16.01 6.9 14.67 7.3 16.09 7.2 Arg 7.64 3.8 12.58 4.9 9.61 4.2 8.22 4.1 9.71 4.3 Gly 22.85 11.3 29.64 11.4 26.99 11.7 25.14 12.4 27.61 12.3 Asp 16.77 8.3 20.77 8.0 18.74 8.1 16.82 8.3 18.76 8.4 MetSO2 2.71 1.3 3.95 1.5 3.21 1.4 2.67 1.3 3.18 1.4 Glu 20.01 9.9 25.17 9.7 22.49 9.7 19.29 9.5 23.68 10.6 Thr 8.29 4.1 10.74 4.1 9.29 4.0 7.62 3.8 9.26 4.1 Ala 10.50 5.2 13.39 5.2 11.88 5.1 9.58 4.7 12.18 5.4 Pro 15.91 7.8 18.30 7.1 16.93 7.3 15.50 7.7 17.37 7.8 Lys 19.45 9.6 19.91 7.7 20.28 8.8 19.08 9.4 21.96 9.8 Tyr 10.34 5.1 14.05 5.4 11.42 4.9 8.56 4.2 12.23 5.5 Val 12.85 6.3 15.89 6.1 14.59 6.3 12.48 6.2 15.08 6.7 Ile 7.98 3.9 11.51 4.4 9.97 4.3 8.52 4.2 10.11 4.5 Leu 13.21 6.5 19.02 7.3 16.02 6.9 13.27 6.6 15.83 7.1 Phe 8.88 4.4 13.31 5.1 10.91 4.7 9.04 4.5 10.79 4.8 Total 203.54 259.06 231.14 202.19 223.82 The amino acids in the table are listed in the order; cysteic acid, histidine, serine, arginine, glycine, aspartic acid, methionine sulphate, glutamic acid, threonine, alanine, proline, lysine, tyrosine, valine, isoleucine, leucine, and phenylalanine. The highlighted amino acids in the indicate those essential amino acids that require supplementation from typical plant-based protein source in fish feed. The totals are a summation of all the individual amino acid quantities in mg/g dry weight for each isolate. The g/100gN column expresses the amino acid equivalence per 100g of protein which is a more conventional way of comparing the amino acid quality in our Pythium biomass to the protein content of other samples in literature. 39

CHAPTER IV: CONCLUSIONS AND FUTURE DIRECTIONS

The promotion of the benefits of omega-3 fatty acids in the overall human physiological and cognitive health (Strobel et al., 2012) has driven higher consumption of fish and other seafoods

(USDA. 2020). Aquaculture has replaced wild-caught fish as a key protein source in global food security while some wild-caught stocks has declined due to overfishing and world-wide stocks have plateaued (Pelletier et al., 2018). Currently, fish meal remains a major source of proteins and primary source of fish oil for farmed fish, yet its costly and unsustainable (Montoya et al.,

2018). This presents new challenges in attaining sustainable fish feed resources in place of fish meal (Estruch et al., 2018). Fish meal contains high levels of protein and nutrient requirements vary across fish species such that a single terrestrial source may not be complete.

Insect meal is now marketed as a natural replacement of fish meal (Nogales et al., 2019).

Insects have high protein and lipid levels even though these contents vary from insect to insect species and depend on their developmental stage and their diet (Nogales et al., 2019).

Carnivorous and omnivorous fish already naturally prey on some insects as part of their diet

(Nogales et al., 2019). Livestock and poultry by products such as meat, viscera, bone, skin, feet, feather and blood are high protein sources and have been employed as protein sources in fish feed (Montoya et al., 2018). Soldier fly and housefly-larvae meal has been researched as a potential fish feed, but it has a poor amino acid profile and low general protein content (Pelletier et al., 2018).

Plant-based proteins have been incorporated as replacement sources of proteins for fish feed.

Some of the plant-based proteins come from soybeans, cotton seeds, sunflower seed, corn, rapeseed and wheat (Montoya et al., 2018). These plant-based protein sources in fish feed boost growth on a similar scale to fish feed (Montoya et al., 2018), however, these protein sources have 40 low levels of essential amino acids such as lysine and methionine and therefore usually require supplementation (Jobling, 2016).

The type of feed fed to fish has an influence on the overall growth rate and immunity.

Consumption of the required amounts of amino acids is especially critical otherwise a deficiency leads to low immunity in fish. For example; low lysine intake causes the dorsal fin to erode, while lack of enough tryptophan, leucine, lysine arginine or histidine causes spinal developmental abnormalities (Pelletier et al., 2018). Individual amino acids are crucial in fish immune defense because they are involved in antibody synthesis and immune regulatory pathways (Egerton et al., 2018). Fish immunity is also related directly to the variety of gut microbes, which changes depending on the diet the fish are fed on (Egerton et al., 2018). Some plant ingredients have been associated with changes in they type of gut microbiota for example a fish feed from soybean meal favors more gram-negative bacteria in the gut while fish meal promotes colonization from mostly gram-positive bacteria (Egerton et al., 2018).

Our study reports that soybean seed-coats are a viable carbon source for the growth of

Pythium in the biosynthesis of omega fatty acids. Pythium secretes cellulose digesting enzymes that are able to digest the fibrous carbohydrates present in the soybean seed coats to release the carbon required for their growth. Overall lipid content of LC PUFAs is low when Pythium is supplied with soybean seed coats as the only carbon source. Supplementation of the cultures with

4% sucrose relieved Pythium of the carbon stress which in turn increased the amount of biomass and omega fatty acids produced by the Pythium isolates. Other carbon sources like corn syrup or glycerol might be better carbon source supplements for the Pythium isolates but they have not been evaluated yet and that is a potential direction we are looking into. 41

Pathogenic Pythium species isolated from the environment are good omega fatty acid producers, however, cultural growth conditions do affect the total amount of omega fatty acids produced. Temperature shift from 250C to 160C during the growth duration of the Pythium cultures does affect the total fatty acid content produced and the resulting total biomass as well.

Higher temperatures (250C) promote faster biomass production while cooler temperatures (160C) promote synthesis of LC PUFAs. Moisture content during the growth process is crucial and therefore we bubbled 500ml/min of oxygen into the culture flasks by bubbling it through autoclaved distilled water. Aeration of samples while growing without supplementation with sucrose does not seem to have an effect in the amount of omega fatty acids produced relative to the total amount of biomass but aeration of the Pythium cultures in concert with 4% supplementation from sucrose boosted omega fatty acids production even more.

Our Pythium biomass has a rich amino acid profile with the essential amino acids known to be limited in plant-based sources in the preferred ranges for fish feed. Further optimization of protein and FA levels relative to the total amount of oomycete biomass may be possible by optimizing inorganic N sources for the Pythium isolates. Regretfully due to the present circumstances of Covid-19, this was not possible. Table 9 below shows a comparison between the amino acid profile of our Pythium biomass with that of protein rich diet; fish meal and soybean meal.

Table 9. Comparison of the amino acid profiles of fish meal, soybean meal and our Pythium biomass

WC001 WC003 WC004 WC005 WC007 Fish Soybean meal meal g/100g g/100g g/100g N g/100g N g/100g N g/100g N g/100g N AA N N Cya 2.0 1.7 2.0 2.0 2.0 His 3.7 3.5 3.6 3.8 3.7 Ser 7.1 6.7 6.9 7.3 7.2 Arg 3.8 4.9 4.2 4.1 4.3 5.86 3.66 Gly 11.3 11.4 11.7 12.4 12.3 Asp 8.3 8.0 8.1 8.3 8.4 MetSO 1.3 1.5 1.4 1.3 1.4 2.30 0.92 2 Glu 9.9 9.7 9.7 9.5 10.6 Thr 4.1 4.1 4.0 3.8 4.1 3.55 1.98 Ala 5.2 5.2 5.1 4.7 5.4 Pro 7.8 7.1 7.3 7.7 7.8 Lys 9.6 7.7 8.8 9.4 9.8 6.01 3.45 Tyr 5.1 5.4 4.9 4.2 5.5 Val 6.3 6.1 6.3 6.2 6.7 Ile 3.9 4.4 4.3 4.2 4.5 Leu 6.5 7.3 6.9 6.6 7.1 Phe 4.4 5.1 4.7 4.5 4.8

The fish meal and soybean meal amino acid values come from a paper by Estruch et al (2018) that analyzed the amino acid profile of various fish feed ingredients. When these values are compared; the amino acid profile for isolate WC007 which is currently our best isolate, is a more balanced protein source than that of soybean meal. Lysine and threonine are high in our

Pythium biomass as compared to that of fish meal. Arginine and methionine values are lower than that of fish meal but can be optimizated with some supplementation of these amino acids.

Further optimization of carbon and nitrogen sources may result in further improvements in the total protein content but are less likely to affect the relative abundance of these amino acids.

The dependability and reproducibility of our protocols is very high. The derivatization protocol curated in our lab is 95% efficient give and take for normal experimental errors and software baseline calculation deviations. Additional experiments that involved independently 43 derivatizing the same samples three days apart achieved the same results for the areas generated by the chromatogram at HPLC which shows that the protocol is highly reproducible. By running additional experiments, we were able to figure out the sufficient amounts of derivatizing reagent to be-750μl of DBAN and 100μl of TEA at 650C for 40 minutes for complete reaction with the complex fatty acid mixture in our 200μl aliquot sample.

This is still a preliminary study. Critical issues that still need to be addressed include the identification of the unknown and co-eluted fatty acid peaks in our fatty acid profile. LCMS is the appropriate tool to address this issue. Our preliminary studies indicate that there is significant variation in omega-3 production by different Pythium isolates. We may not yet have identified the best Pythium strains for industrial production. Finally, pilot scale production of our oomycete biomass is needed to test its palatability an ingredient in fish food, and its overall effects on fish health.

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