EXPLORATION AND OPTIMIZATION OF THREE ENZYMATIC ROUTES TO CONVERT LINOLEIC ACID INTO N-HEXANAL

By

ERICA AMATO SIMMONS

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2017

© 2017 Erica Simmons

To my loving parents, Pasquale and Denise Amato and husband, Aaron Simmons

ACKNOWLEDGMENTS

I would first like to thank Dr. Jon Stewart for his continuing support and encouragement throughout my Ph.D. work. His guidance has been essential, especially at times when I just wanted to quit the project due to constant failures. He motivated me to keep going and to see the project through to completion, without which I would have never been able to ultimately find a method to convert linoleic acid into n-hexanal. I also need to mention the contributions from other faculty members such as Dr. Horenstein, Dr. Butcher and Dr. Bruner for always being available for advice regarding my project and for allowing me to use lab equipment when it was broken in my laboratory. Much of the reaction progress mentioned throughout this work would not have been possible without the help of their labs. I need to specially thank my undergraduate mentee, Olivia Toudjarov, who despite her hectic schedule would come to the laboratory to do research. I would also like to thank the various members of my research group for always being willing to help when asked.

Next I would like to thank the people who got me through this graduate school program including Mayra Rostagno, Jessica Cash, Sandy Guntaka, and Kim Stewart. They were always willing to give me suggestions when I didn’t know where to go next in my research. In addition, they were crucial in making sure I didn’t quit my project when it seemed impossible to finish.

They encouraged me to try other directions and to take breaks to clear my mind before returning to the project. It is always encouraging to know that various disciplines can come together to approach a project from a new perspective and think of new strategies despite the fact that it is not their strength.

Finally, I would like to thank my family for constant encouragement and support when hard times came up. Being over a thousand miles away from them was not easy throughout this

Ph.D. program, but we always found ways of seeing each other and were in constant

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communication. At times, they were the motivation I needed to go into lab the next day and push through the constant failures that accumulated. I need to give a special thanks to my husband,

Aaron Simmons, for putting up with me when my research stressed me out and became frustrating. He managed to always find ways of making me forget about work and to relax. His love and support always push me to be a better person and ultimately to complete this Ph.D. program.

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TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 10

LIST OF FIGURES ...... 11

ABSTRACT ...... 15

CHAPTER

1 INTRODUCTION ...... 16

Short Chain Aldehydes and Ketones ...... 16 European Union Regulations on Natural Flavors ...... 18 Current n-Hexanal Synthetic Routes ...... 18 Chemical Synthesis Approaches ...... 18 Natural Sources ...... 20 Bioprocesses ...... 22 New Strategies for Producing n-Hexanal from Linoleic Acid ...... 24 Castor Oil Pyrolysis ...... 26 Carotenoid Cleavage Dioxygenases ...... 27 Identification and Characterization ...... 27 Carotenoid cleavage dioxygenase 1 (EC: 1.14.99.n4) ...... 29 Carotenoid cleavage dioxygenases 7 (EC:1.13.11.68) and 8 (EC:1.13.11.691) ...... 31 CCD Enzyme Structure ...... 33 Mechanism ...... 36 Mutagenesis Studies ...... 39 Applications of CCD1 ...... 39 Lipoxygenases ...... 41 Identification and Characterization ...... 41 Enzyme Structure ...... 43 Mechanism ...... 46 Lineolate 13-Hydratase ...... 48 Identification and Characterization ...... 48 Enzyme Structure ...... 51 Mechanism ...... 54

2 USING CAROTENOID CLEAVAGE DIOXYGENASES TO CLEAVE LINOLEIC ACID INTO N-HEXANAL ...... 56

Background ...... 56 Experimental Strategy ...... 56 n-Hexanal Quantitation ...... 56 CCD Production and Isolation ...... 57

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Cleaving Linoleic Acid to n-Hexanal Using CCD1 Enzymes ...... 58 Experimental Procedures ...... 58 Light Scattering Experiments ...... 58 n-Hexanal Quantitation ...... 59 1. Solid phase micro extraction (SPME) ...... 59 GC/MS analysis using EDA2_MTH ...... 59 2. 2,4-Dinitrophenylhydrazine (DNPH) derivitization ...... 60 3. Solvent extraction using d12-n-hexanal ...... 61 Resin recoveries ...... 61 Linoleic Acid Quantitation ...... 61 FAME derivitization ...... 61 MSTFA derivitization ...... 62 GC/MS analysis: EA1_METH ...... 62 ZmCCD1 Expression ...... 63 Construction of pEA1 ...... 63 Construction of pEA2 ...... 64 Construction of pEA3 ...... 64 AtCCD1 Expression ...... 64 Acetylation of AtCCD1 ...... 65 CLEA Preparation ...... 65 β-Apo-8’-Carotenal Oxidations ...... 66 HPLC analysis: EA_MTH1 ...... 67 Protein Purification ...... 67 Gluathione S-transferase affinity purification ...... 67 Regeneration of GST column ...... 68 Amylose affinity purification ...... 68 Regeneration of amylose column ...... 68 Sephacryl S-200 gel filtration chromatography ...... 68 Bradford and bicinchoninic assays ...... 69 Enzyme Reactions ...... 69 ZmCCD1 reactions ...... 69 Linoleic acid solution ...... 58 AtCCD1 reactions ...... 69 Autoxidation Trials ...... 70 Biphasic reactions ...... 70 GC/MS analysis: JON_METH ...... 70 Site Saturation Mutagenesis95 ...... 71 Results and Discussion ...... 72 Linoleic Acid Behavior under Aqueous Conditions- Light Scattering Studies ...... 72 n-Hexanal Quantitation ...... 75 SPME quantitation ...... 75 Other methods for extracting and detecting n-hexanal ...... 76 Linoleic Acid Quantitation ...... 78 Cloning and Expression of AtCCD1 ...... 79 MBP-AtCCD1 Cleavage of Linoleic Acid ...... 82

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Effects of pH, temperature and buffer on linoleic acid cleavage into n-hexanal by MBP-AtCCD1 ...... 83 Effects of CLEA and acetylation on the MBP-AtCCD1 cleavage activity of linoleic acid ...... 84 Purification of MBP-AtCCD1 for enzymatic reactions ...... 85 Effects of alcohol in the reaction mixtures to solubilize linoleic acid ...... 86 Autoxidation of Linoleic Acid ...... 87 Other AtCCD1 Substrates ...... 89 Ligand Binding Studies of MBP-AtCCD1 ...... 90 Homology Model of AtCCD1 ...... 93 AtCCD1 Mutagenesis studies ...... 96 Conclusion ...... 98

3 PRODUCTION OF 13S-HYDROPEROXY-9Z,11E-OCTADECADIENOIC ACID USING LIPOXYGENASE (LOX1) ISOLATED FROM SOYBEANS ...... 99

Background ...... 99 Experimental Strategy ...... 100 13S-Hydroperoxy-9Z,11E-Octadecadienoic Acid Cracking ...... 100 Lipoxygenase ...... 100 Linoleic Acid Conversion and Reduction ...... 100 Experimental Methods ...... 101 Pyrolysis of Castor Oil ...... 101 GC/MS analysis: JON_METH ...... 101 Lipoxygenase Reactions ...... 102 13-(S)-Hydroperoxy-9Z,11E-Octadecadienoic Acid Quantitation ...... 102 1. Ferrous-xylenol orange (FOX) method ...... 102 2. UV-Vis ...... 103 3. FAME/MSTFA derivitization ...... 103 Lipoxygenase Isolation from Soybeans ...... 103 13S-Hydroperoxy-9Z,11E-Octadecadienoic Acid Cracking ...... 104 Results and Discussion ...... 104 Castor Oil Pyrolysis ...... 104 Biotransformation of Linoleic Acid Using Lipoxygenase ...... 105 Optimization of the Lipoxygenase Reaction with Linoleic Acid ...... 107 Effects of the type of lipoxygenase used in the biotransformations ...... 107 Effects of TCEP in the reaction mixtures ...... 108 Effects of increasing the ethanol concentration in the reaction mixtures...... 109 Other Substrates Tested for Lipoxygenase Activity ...... 109 Thermal Cracking of 13S-Hydroperoxy-9Z,11E-Octadecadienoic Acid ...... 110 Conclusion ...... 112

4 PRODUCTION OF 13S-HYDROXY-9(Z)-OCTADECENOIC ACID USING LINEOLATE 13-HYDRATASE FROM L. ACIDOPHILUS ...... 113

Background ...... 113 Experimental Procedures ...... 114

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Cloning of pEA4 ...... 114 Cloning of pEA6 ...... 114 Lineolate 13-Hydratase Expression ...... 115 Bicinchoninic Acid Assay ...... 115 Lineolate 13-Hydratase Enzymatic Reactions ...... 115 13-Hydroxy-9Z-Octadecenoic Acid Cracking Procedure ...... 116 GC/MS analysis: EA1_METH ...... 116 Results and Discussion ...... 117 Cloning and Expression of Lineolate 13-Hydratase ...... 117 Detection of 13S-Hydroxy-9Z-Octadecenoic Acid ...... 121 Hydration of Linoleic Acid with Crude Lysate Suspensions ...... 121 Hydration of Linoleic Acid with Whole Cells ...... 123 Various Buffer Systems in the Hydratase Reactions ...... 125 Effects of Temperature on the Hydration of Linoleic Acid ...... 126 Multiple Enzyme Additions to the Hydratase Reactions ...... 128 Time Course Study for the Hydratase Reactions ...... 129 Addition of Detergent in the Reaction Mixtures ...... 130 Extraction Methods to Remove Biotransformation Products ...... 130 Cracking of Biotransformation Products from Lineolate 13-Hydratase Reactions ...... 133 Scaling Up Lineolate 13-Hydratase Reactions ...... 137 Other Substrates Evaluated with Lineolate 13-Hydratase ...... 138 Future Directions ...... 138 Conclusion ...... 139

APPENDIX

A ADDITIONAL PLASMID INFORMATION ...... 141

Nucleic Acid Sequences ...... 141 Plasmid Maps...... 144 Primer Sequences ...... 146

B FATTY ACID STRUCTURES ...... 147

C ADDITIONAL GC/MS DATA ...... 149

GC Chromatograms ...... 149 Mass Spectra ...... 150

D NMR DATA ...... 152

LIST OF REFERENCES ...... 153

BIOGRAPHICAL SKETCH ...... 160

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LIST OF TABLES

Table page

1-1 Volatile natural flavor compounds...... 16

1-2 US versus EU regulations for natural flavorings ...... 18

1-3 Key differences between the isozymes of soybean LOX enzymes ...... 43

2-1 Conditioning parameters for each of the SPME fibers evaluated ...... 59

2-3 Inhibition trial results for various materials evaluated with MBP-AtCCD1 ...... 91

4-1 Whole cell-mediated hydrations of linoleic acid at two temperatures ...... 126

4-2 Crude lysate-mediated hydrations of linoleic acid at two temperatures ...... 127

4-3 Six hour whole cell-mediated hydrations of linoleic acid at two temperatures ...... 127

A-1 List of Sequence Altering Primers for AtCCD1 ...... 146

A-2 List of Mutagenic Primers for AtCCD1...... 146

B-1 Significant chemical structures ...... 147

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LIST OF FIGURES

Figure page

1-1 Synthesis of n-hexanal by utilizing the Grignard synthetic route ...... 19

1-2 Synthesis of n-hexanal by utilizing the Stephen reaction route ...... 19

1-3 Synthesis of n-hexanal from linoleic acid through ozonolysis ...... 20

1-4 Pyrolysis of ricinoleic acid into n-heptanal and undecylenic acid ...... 22

1-5 n-Hexanal production from lipoxygenase and hydroperoxide lyase ...... 24

1-6 The three proposed routes of converting linoleic acid into n-hexanal ...... 25

1-7 Various carotenoid examples ...... 28

1-8 Some examples of apocarotenoids ...... 28

1-9 Cleavage of apocarotenoid β-apo-10’-carotenal into retinal and a di-aldehyde ...... 29

1-10 Synthetic substrate β-apo-8’-carotenal ...... 29

1-11 Cleavage of β-carotene at the 9/10 position ...... 30

1-12 Carotenoids accepted by CCD1 and their cleavage positions ...... 31

1-13 Apo-10’-lycopenal being cleaved by OsCCD1 at the 7/8 alkene ...... 31

1-14 Sequential cleavage of CCD7 and CCD8 to produce carlactone ...... 32

1-15 The octahedrally coordinated Fe in the active site of the ACO enzyme ...... 33

1-16 Crystal structure of ACO ...... 34

1-17 Hydrophobic surface of ACO ...... 35

1-18 Homology model of ZmCCD1 ...... 36

1-19 Proposed pathways for carotenoid cleavage by CCD enzymes ...... 37

1-20 Several substrates known to be cleaved by AtCCD1 ...... 40

1-21 Linoleic acid aligned with two known substrates of CCD1 ...... 41

1-22 Cis, cis-1,4-pentadiene systems ...... 42

1-23 1.4Å crystal structure of LOX-1 ...... 44

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1-24 Octahedrally coordinated iron in the soybean LOX1 active site ...... 45

1-25 Active site depiction of soybean LOX-3 harboring linoleic acid ...... 46

1-26 The 13-LOX enzyme mechanism and its regioselectivity using linoleic acid ...... 47

1-27 The three proposed radical rearrangement mechanisms for the 13-LOX ...... 48

1-28 Hydration of linoleic acid to form the 13- and 10- hydroxyl products ...... 49

1-29 Crystal structure of a fatty acid double bond hydratase from L. acidophilus ...... 52

1-30 Hydrophobicity plot of an LAH ...... 52

1-31 Crystal structure of Oleate hydratase from Elizabethkindia meningoseptica ...... 53

1-32 Active site of Oleate hydratase with FAD ...... 54

1-33 The proposed mechanism for alkene hydration adapted from Engelder et al...... 55

2-1 Predicted linoleic acid cleavage position of CCD1 into n-hexanal ...... 56

2-2 DNPH derivitization of hexanal...... 60

2-4 Light scattering results mimicking actual enzymatic reaction conditions ...... 74

2-6 GC of d12-n-hexanal and n-hexanal separation ...... 78

2-7 Construction of pEA1 from pBS2 and pUC57-AtCCD1 ...... 79

2-8 10% SDS-PAGE of the purified GST-AtCCD1 fusion protein ...... 80

2-9 The construction of pEA2 from pMAL-c5x and pEA1 ...... 81

2-10 Expression level comparison ...... 82

2-11 n-Hexanal production from enzymatic and non-enzymatic reactions ...... 83

2-12 Comparison of various changes to MBP-AtCCD1 reaction parameters ...... 84

2-13 Purification of AtCCD1-MBP with amylose column ...... 85

2-14 Comparison of MBP-AtCCD1 purification ...... 86

2-15 Effects of increasing the concentration of EtOH in the reaction systems...... 87

2-16 Autoxidation of linoleic acid compared to enzymatic reactions ...... 88

2-17 Effects of adding catalase to the reaction systems ...... 89

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2-18 HPLC data for β-apo-8’-carotenal reactions ...... 91

2-19 Homology model of AtCCD1 with bound substrate predicted by I-TASSER ...... 94

2-20 Catalytic Fe coordination within the active site tunnel of AtCCD1 ...... 95

2-21 Overlap of 3-hydroxy-8’-carotenal with linoleic acid ...... 95

2-22 Residues respoonsible for positioning linoleic acid near the catalytic Fe ...... 96

2-23 The two positions proposed for mutagenesis studies ...... 97

3-1 Experimental strategy of using lipoxygenase to create n-hexanal ...... 99

3-2 Castor Oil Pyrolysis set up ...... 101

3-3 Microdistillation flask used for cracking procedure and associated condenser ...... 104

3-4 Fatty Acid Structures ...... 105

3-5 UV-Vis spectrum for lipoxygenase reactions ...... 105

3-6 Mass spectrum analysis of 13-(S)-hydroxyoctadeca-9Z, 11E-dienoic acid ...... 106

3-7 Comparison of lipoxygenase conversion of 1 mM linoleic acid ...... 107

3-8 Comparison of reactions with TCEP included and those without ...... 108

3-9 GC difference chromatogram of the cracking products from lipoxygenase ...... 110

4-1 Comparison of the cracking substrates evaluated ...... 113

4-2 Construction of the pEA4 plasmid from pET-15b and pUC57 ...... 117

4-3 Comparison of the soluble protein present in pEA4 overexpression systems ...... 119

4-4 Construction of the pEA6 plasmid from pACYC Duet-1 and pEA4 ...... 120

4-5 Soluble proteins present in pEA6 18 hour overexpression ...... 120

4-6 Mass spectrum analysis of 13-hydroxy-9Z-octadecenoic acid ...... 121

4-7 Determination of the optimal crude lysate concentration for hydratase reactions ...... 122

4-8 Determination of optimal linoleic acid concentation for crude lysate reactions ...... 123

4-9 Determination of the optimal linoleic acid concentration for whole cell reactions ...... 124

4-10 Buffer system comparison of 10 mM linoleic acid conversion ...... 126

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4-11 Multiple crude lysate additions to the hydratase reaction ...... 128

4-12 Time course study for the hydration of 50 mM linoleic acid ...... 129

4-13 Continuous extraction apparatus using heat ...... 131

4-14 Continuous extraction set up using an HPLC pump and EtOAc ...... 132

4-15 Continuous extraction layer separation with an HPLC pump ...... 133

4-16 Chromatogram of cracking products dissolved in peanut oil ...... 134

4-17 Chromatogram of carrier-free hydratase product cracking ...... 135

4-18 3-neck flask cracking apparatus ...... 136

4-19 Chromatogram of 3-neck flask cracking products ...... 137

A-1 Sequence of Zea mays CCD1 ...... 141

A-2 Sequence of Arabidopsis thaliana CCD1 ...... 142

A-3 Sequence of Glycine max Lipoxygenase 1 ...... 143

A-4 Sequence of Lactobacillus acidophilus strain KCTC Lineolate 13-Hydratase ...... 143

A-5 Plasmid map of pEA1 ...... 144

A-6 Plasmid map of pEA2 ...... 145

A-7 Plasmid map of pEA3 ...... 145

A-8 Plasmid map of pEA4 ...... 145

A-9 Plasmid map of pEA6 ...... 146

C-1 GC chromatogram for lipoxygenase biotransformation products ...... 149

C-2 GC chromatogram for lineolate 13-hydratase biotransformation products ...... 149

C-3 Mass spectrum analysis of linolenic acid hydration with lineolate 13-hydratase ...... 150

C-4 Mass spectrum analysis of palmitoleic acid hydration ...... 150

C-5 Mass spectrum analysis of methyl lineolate hydration ...... 151

D-1 1H NMR of DNPH derivitized n-hexanal ...... 152

D-2 1H NMR of pure n-hexanal ...... 152

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Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

EXPLORATION AND OPTIMIZATION OF THREE ENZYMATIC ROUTES TO CONVERT LINOLEIC ACID INTO N-HEXANAL

By

Erica Amato Simmons December 2017

Chair: Jon D. Stewart Major: Chemistry

Three distinct pathways were explored for the conversion of linoleic acid into n-hexanal through an all-natural route. The first utilized a carotenoid cleavage dioxygenase enzyme that ultimately resulted in an insufficient amount of n-hexanal production (~0.05 mg, 10% yield).

This was attractive due to it being a single step and using only molecular oxygen as a cofactor.

Several optimization trials were attempted; however, this was the maximum n-hexanal quantity reached. The second route relied on a lipoxygenase enzyme to put a peroxide group on the C13 of linoleic acid and left both alkene bonds on the carbon chain intact. The peroxide group was reduced to an alcohol prior to undergoing thermal cracking to produce n-hexanal. When undergoing cracking, the hydrated linoleic acid underwent a rearrangement and could not form the amounts n-hexanal necessary for industrial purposes (~0.16 mg, 0.04% yield). This rearrangement product, 2-pentyl furan, could not yield n-hexanal. Finally, the third route utilized lineolate 13-hydratase to directly hydrate the C13 of linoleic acid and removed the 12, 13 alkene, making 13-hydroxy-9Z-octadecenoic acid. The same thermal cracking procedure was utilized for this substrate and produced significant amounts of n-hexanal (~39 mg, 1% yield). In addition, there were fewer side products observed in the cracking product mixture, making n-hexanal isolation and purification simpler.

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CHAPTER 1 INTRODUCTION

Short Chain Aldehydes and Ketones

Short chain aldehyde and ketone molecules are significant in the fragrance and flavoring industries, primarily due to their strong, naturally occurring aromas. A few examples include n-hexanal, n-nonanal, and cinnamaldehyde.1 Several common aldehyde and ketone compounds used for these purposes and their associated scents are listed in Table 1-1. In some cases, molecules have been found to change their scent depending on their absolute configurations. Carvone is the most familiar example. The scent of (R)-carvone resembles caraway, while (S)-carvone resembles spearmint.2

Table 1-1. Volatile natural flavor compounds

Compound Structure Associated Scent n-Hexanal Fruity1 n-Octanal Citrus1 n-Nonanal Floral1 n-Decanal ‘Fatty’ odor 1

Cinnamaldehyde Cinnamon1

(S)-Carvone Spearmint3

Some aldehyde compounds also have antimicrobial activities. It is thought that they play a role in plant defense against pest and pathogen attack.4,5 For example, Lanciotti et al. determined that n-hexanal significantly extended the shelf life of apple slices.5 They evaluated microbial growth at 4°C and 15°C in the presence of n-hexanal and found that at 4°C, a total

16

inhibition of mesophilic bacteria occurred. In addition, at 15°C n-hexanal inhibited mold, yeast, mesophilic, and psychrotrophic bacteria formation on the apple slices. The antimicrobial activity of aldehyde compounds is of industrial interest for its ability to increase the shelf life of various fruits and vegetables and potentially eliminating the possibility of food-borne pathogens. In addition, there is an increasing demand for minimally processed foods and these aldehyde compounds have the potential to help reach this goal. Lastly, short chain aldehyde and ketone molecules can act as messengers in communication between themselves and other organisms.6

Our industrial partners are interested in synthesizing n-hexanal from linoleic acid while following the European Union (EU) regulations governing natural flavors. n-Hexanal is considered one of the ‘green’ note compounds due to its characteristic fresh green odor of cut leaves and is of particular importance in the fragrance and flavor industries.7 It is estimated that the worldwide use of green note compounds is $20-40 million per year.8

Aldehyde compounds are generally volatile and have low solubility in aqueous solutions as their molecular weights increase.9 They are also highly reactive, which increases the difficulties in working with this class of compounds. Moreover, their volatility makes them difficult to quantitate and analyze. They are also easily oxidized and can react with various oxidizers including molecular oxygen to form the corresponding carboxylic acids.9 For example, over the course of 24 hours, it was found that 95% of n-hexanal autoxidized into various compounds, primarily hexanoic acid.10 In addition, it was found that once decomposed, the by-products could recombine to form compounds with molecular weights exceeding that of n-hexanal. The dominant autoxidation products included esters and lactones, with carbonyls, acids, alcohols, and hydrocarbons being present in smaller numbers.10 Autoxidation likely

17

occurs via a free radical mechanism.11 Finally, it is known that saturated aldehydes undergo polymerization under the influence of oxygen and heat and that these polymers further react to form the high molecular weight compounds observed from autoxidation.10

European Union Regulations on Natural Flavors

In general, the EU regulations are comparable to those in the United States (Table 1-2).12

Other global regulations are usually based on one of these two standards when determining whether a flavor is natural. The EU regulations are slightly stricter because the European Food

Safety Authority (EFSA) is more restrictive with respect to permitted processes and a list of these processes can be found in EU directive 1334/2008, which is much more lengthy than in its counterpart in the US.

Table 1-2. US versus EU regulations for natural flavorings12

US EU Production methods are vaguely defined Must be produced by an approved method Specific biological sources are defined in the Biological sources are vaguely defined 21CFR regulations No corresponding definition Natural flavoring substances are naturally present or have been identified in nature *Adapted from Sigma

The EU regulations were put in place in 2009 and an approved list of natural flavorings was created in 2012.13 They lay out general requirements for the safe use and definitions for different types of flavorings. While there are several compounds on the list of approved natural flavors, n-hexanal does not appear. To the best of our knowledge there are few aldehyde compounds on this list.

Current n-Hexanal Synthetic Routes

Chemical Synthesis Approaches

The majority of methods currently used to synthesize flavor aldehyde and ketone molecules are based on traditional chemical synthesis. These methodologies, however, are

18

proving to be increasingly ineffectual in the flavoring industry due to increasing Food and Drug

Administration (FDA) regulations. Figures 1-1 and 1-2 outline two of the more common synthetic routes for industrial purposes: the Grignard and Stephen reactions. For the fragrance industry, these synthetic routes are acceptable processes. This is not the case for the flavoring and food industries that need to be able to make these compounds without the use of potentially harmful materials.

Figure 1-1. Synthesis of n-hexanal by utilizing the Grignard synthetic route14

Figure 1-2. Synthesis of n-hexanal by utilizing the Stephen reaction route14

In addition to Figures 1-1 and 1-2, flavor aldehyde and ketone molecules can also be prepared from alkenes using ozonolysis (Figure 1-3) or by using heavy metal catalysts such as chromium (Cr), osmium (Os) and ruthenium (Ru).15 These methods, however, use harsh conditions, are expensive and have mediocre yields and selectivities. For these reasons, the

Grignard and Stephen reactions are the primary routes to synthesizing short chain flavor aldehydes and ketones.

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Figure 1-3. Synthesis of n-hexanal from linoleic acid through ozonolysis16

Natural Sources

n-Hexanal can be isolated directly from various plant tissues (specifically leaves) under specific physiological conditions that favor catabolic reactions at low levels.6 Volatile aldehydes can also be found in trace quantities in essential oils (<10%, with a majority present at <1%).

These volatile aldehydes are obtained from plants through distillation or by extraction with various organic solvents. An industrial undertaking, however, can be an expensive operation with such low quantities of the target aldehyde per plant mass.6

Often, plant sources for desired volatile aldehydes are slow-growing and difficult to find.

It was therefore hypothesized that in vitro cultured plants would speed up the biomass propagation rate by growing them under controlled conditions so that the metabolites would be immediately available.6 The actual production rate for the metabolites were in the range of 0.1-

0.01 g/L/day, which was orders of magnitude lower than expected and also lower than that from the intact plants.17 A process based on this strategy would require very large biomass production.

With high cell titers (larger than 15-20 g/L), growth inhibition can occur. While there are strategies to overcome the growth inhibition, there is still the viscosity problem that is difficult to deal with, which leads to reduced mixing and oxygen supply.

It was discovered that the production of volatile aldehydes could be induced in cultured plant tissues and cells by a variety of chemical and physical factors.6 This route, however, influenced the essential oil yield that would eventually provide the volatile aldehydes. While

20

treatments were found that increased the essential oil yield by 5-fold, it was still less than that achieved by the intact plants.18, 19 Based on these results, the use of cultured plant tissues for volatile aldehyde production does not seem to be promising.

Volatile aldehyde and alcohol compounds can also be produced by cultured microorganisms.6 In general, bacteria and fungi are sturdier than the plant biomass used in the previously mentioned methods. They can withstand frictional stress from shaking and endure various pH, temperature, and salt conditions. There are several reviews covering the vast array of microorganisms that produce these volatiles.20,21,22 For example, benzaldehyde can be produced by cultured Goetrichum candidum with a yield of 1.6 g/L. This fungus can also produce 9.5 g/L of 2-hexanoic acid ethyl ester in the same culture.23 Together these compounds produced a fruity fragrance. There are numerous other examples of microorganisms producing volatile aldehyde and alcohol compounds, however, it is still economically favorable to produce these compounds by chemical synthesis that provide higher yields.

Another natural source of n-hexanal comes from the autoxidation of lipids, specifically fatty acids. This autoxidation occurs via a free radical mechanism that is initiated by the abstraction of hydrogen from the methylene carbon of an unsaturated fatty acid.24 This autoxidation can be induced by a variety of conditions, including increased oxygen availability, or by the presence of free radical compounds such as iron and ascorbic acid. The primary products formed in these reactions are aldehydes by scission of the fatty acids on either side of the radical.24

Finally, in the late 1980s, an experiment was performed called castor oil pyrolysis. In this process, neat castor oil in the presence of rosin was heated to 400°C with a bare flame to produce large quantities of n-heptanal and undecylenic acid via what is believed to be a free radical

21

process.25 Castor oil contains 85-95% ricinoleic acid (Figure 1-4) in its triglyceride form. As mentioned previously in Table 1-1, n-heptanal is also a flavor aldehyde and this process has potential industrial applications due to its high yields, cost efficiency and simplicity. The significance of this process to the current study will be discussed in later sections.

Figure 1-4. Pyrolysis of ricinoleic acid into n-heptanal and undecylenic acid

Bioprocesses

With the growing popularity of natural products in the food industry, biocatalysis has become an important contributor in the past decade. In general, biotechnology reduces the market cost of natural product synthesis through the use of non-polluting methods.6 Bioprocesses have additional advantages including the production of chirally pure compounds at high yields

(rather than racemic mixtures), selective modification of a particular substrate, and functionalizing chemically inert carbons.21 Bioprocesses for the production of volatiles have their limitations, however. Some flavor metabolites such as vanillin lead to feedback inhibition of their biotransformations and if they are not protected in some form they could further react to form side products.26 Another common problem is that the substrates and products may not easily dissolve in an aqueous medium. Methods have been developed to add the substrate continuously to the enzymatic reaction and remove the product as it is being produced.6 Product removal can be accomplished in various ways. Two of the most common include the use of a two-phase system or employing solid organic phases such as resins. The two phase system

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includes an aqueous portion and an organic solvent to extract the desired product. It has been reported that the addition of amberlite XAD-7 to Cinchona ledgeriana cultures increased the production of anthraquinones by 15 times compared to without the resin.27 The addition of charcoal to Ma. Chamomilla suspensions led to 20-60-fold increases in coniferyl aldehyde production.28

Bioprocesses for the synthesis of flavor aldehyde and ketone molecules themselves, however, are significantly limited. Companies need to be able to produce large amounts of these compounds as economically as possible. As stated previously, flavor aldehydes are volatile, reactive, and difficult to quantitate. These qualities make them difficult to work with and more importantly, to isolate as pure products.

In a majority of food process development, it is preferable to have a starting material that can be isolated from natural products. In the case of this study, unsaturated fatty acids are the desirable substrate. This class of compounds can be easily isolated from a variety of oils, depending on the desired fatty acid composition. A majority of these oils are plant based, for example: sunflower seed oil (~75% linoleic acid), olive oil (~80% oleic acid), castor oil (~80% ricinoleic acid) and coconut oil (~52% lauric acid).29 The majority of isolated enzyme-catalyzed bioconversions involve lipid-derived volatile products.6 As mentioned previously, n-hexanal is considered a ‘green’ note volatile and this group is particularly important when it comes to crude enzyme preparations of fatty acids due to their increasing popularity and the need for natural production.

The only current bioprocess for the production of n-hexanal from fatty acids utilizes a 2 step mechanism that comprises soybean lipoxygenase followed by a hydroperoxide lyase (Figure

1-5).

23

Figure 1-5. n-Hexanal production from lipoxygenase and hydroperoxide lyase30, 31

The position 13-selective lipoxygenase enzyme used in this route is generally isolated from soybeans and is both stable and commercially available. It is an efficient, high yielding enzyme with reported yields of up to 80% with a single addition of the lipoxygenase.32, 33 Non- enzymatic peroxidation of these compounds leads to unspecific product mixtures, therefore, the use of lipoxygenase in industrial processes is preferred.34 Unfortunately, the hydroperoxide lyase is an unstable enzyme and none are commercially available. This enzyme has yields of around

54%, albeit at small concentrations.32 Numerous groups are attempting to increase the effectiveness of hydroperoxide lyase, but have found little success to date. We therefore decided to explore other bioprocesses in the production of n-hexanal that would allow it to retain its

“natural” status.

New Strategies for Producing n-Hexanal from Linoleic Acid

We identified three potential strategies for converting linoleic acid into n-hexanal by methods that would meet the EU standards for natural flavors (Figure 1-6). The first was based on a position 13-selective lipoxygenase, which would incorporate an initial C13-hydroperoxy moiety that would then be reduced and cleaved thermally to n-hexanal in a reaction analogous to that of n-heptanal production by castor oil pyrolysis. A second route also involved thermal cleavage of C13-hydroxy derivative of linoleic acid, produced directly by a position 13-selective

24

hydratase. The final route featured direct conversion of linoleic acid to n-hexanal by a carotenoid cleavage dioxygenase (CCD).

Figure 1-6. The three proposed routes of converting linoleic acid into n-hexanal

We first explored the CCD approach since it required only a single step and molecular oxygen as the sole co-substrate. After several attempts, however, none were successful. Linoleic acid is known to be susceptible to autoxidation and there was never definitive proof that the n- hexanal observed had resulted from enzymatic cleavage rather than autoxidation. Control reactions lacking the enzyme yielded only slightly less n-hexanal production compared to the reactions containing enzyme.

Our first attempt at producing n-hexanal by the pyrolysis route used a lipoxygenase enzyme. n-Hexanal was produced, albeit at a low concentration. The primary product was derived from a rearrangement of linoleic acid, which made it impossible to isolate the desired

C13-hydroxy derivative in sufficient quantities.

25

We successfully produced n-hexanal from linoleic acid by pyrolyzing a monounsaturated

C13-hydroxyl derivative, the product of a position 13-selective hydratase. The hydratase product closely resembled the ricinoleic acid structure used in the castor oil pyrolysis. Furthermore, the yield of the hydratase reaction provided sufficient material to carry out the pyrolysis procedure on the neat product. The properties of each of the enzymes required for the three strategies are reviewed in the following chapters.

Castor Oil Pyrolysis

The inspiration for the lipoxygenase and lineolate 13-hydratase routes to n-hexanal synthesis came from castor oil pyrolysis. Castor oil contains 85-95% ricinoleic acid in its triglyceride form. In addition, the rosin used to aide in the pyrolysis procedure is produced from pine trees, therefore following the EU regulations of using all natural components. We reproduced the results of this experiment in our lab and confirmed that it is an efficient process.25

This procedure was attractive due to the high yield of n-heptanal, low cost of the castor oil, simplicity of the procedure, and the use of all natural substrates. To produce n-hexanal, the hydroxyl needs to be at the C13 (rather than at C12 as in ricinoleic acid). We therefore sought methods to make a 13-hydroxy derivative of linoleic acid enzymatically. It was unknown if the presence of the second double bond on linoleic acid would cause problems in the pyrolysis procedure. A lipoxygenase enzyme can be used to hydroxylate the linoleic acid at the 13 carbon position (Figure 1-5). This process conjugates the alkenes and creates an alkyl alcohol. A second route that was discovered later was the use of the enzyme lineolate 13-hydratase. The hydratase leaves a single alkene and an alkyl alcohol on the 13 carbon position (Figure 1-6). This later product is more analogous to the ricinoleic acid in castor oil (Figure 1-4).

26

Carotenoid Cleavage Dioxygenases

Identification and Characterization

Oxidative cleavage of alkenes is widely utilized in synthetic organic chemistry in order to introduce oxygen functionality in molecules, remove protecting groups, and degrade larger molecules.16 As mentioned previously, there are synthetic methods to oxidatively cleave alkenes; however, they can lack regioselectivity and sometimes require harsh conditions and/or reagents.

By contrast, enzymes are able to activate molecular oxygen and catalyze the cleavage of alkenes under physiological conditions with high regioselectivity.

Carotenoid Cleavage Dioxygenases (CCDs) are a large family of non-heme, iron dependent enzymes that selectively cleave one or more double bonds in the polyene structures of carotenoid compounds. Depending on the cleavage position, they can create either aldehyde or ketone molecules from carotenoids.35 These products have important roles in plant growth, protection against light exposure, as antifungal agents, and as sources of fragrances for reproduction.36 There was no direct evidence for the existence of CCD enzymes until the discovery of Zea mays Vp14 (NCED1).37 NCED enzymes differ from CCD due to their involvement in abscisic acid biosynthesis; however, they are a part of the same family of enzymes.38 After Vp14 had been identified, several other CCD enzymes were discovered including β-carotene oxygenase (BCO) in mammals and CCD1, CCD7 and CCD8 in plants.35 A majority of CCD enzymes are membrane associated, with only a few reported to be located in the cytosol.39

The main substrates for CCD enzymes are carotenoids and Figure 1-7 shows a few examples.

27

(a) (b)

(c) Figure 1-7. Various carotenoid examples (a) β-carotene, which exhibits an orange pigment in carrots, (b) violaxanthin, which exhibits an orange pigment in pansies, and (c) astaxanthin, which exhibits a red pigment in salmon and shrimp.

Carotenoids are C40 compounds that are widely distributed in nature and have important metabolic and hormonal functions in prokaryotes, animals, fungi, and algae.40 In higher plants, they mediate fragrance, fruit color, and aroma. Alkene cleavage products of carotenoids are known as apocarotenoids and these regulate plant growth and development as stated previously.40 Figure 1-8 shows some examples of apocarotenoids. Carotenoids have characteristic conjugated double bond systems, which ultimately determine their apparent color.

As the amount of conjugation increases, the color changes from yellow to orange to red. This is the reason that they are commonly used as chromophores.41 Carotenoids also have low water solubility; however, the CCD enzymes do not stand up well to organic solvents. The use of various detergents in these reactions is necessary and they also aide in the release of the CCD enzymes from the membranes.

(a)

(b) Figure 1-8. Some examples of apocarotenoids (a) The products of the CCD1 cleavage of β- carotene.35 (b) The products of the cleavage of β-apo-10’-carotenal

28

Some apocarotenoid compounds can themselves act as substrates for CCDs. For example,

β-apo-10’-carotenal can be further cleaved by CCD enzymes to form retinal and a small di- aldehyde (Figure 1-9).41

Figure 1-9. Cleavage of apocarotenoid β-apo-10’-carotenal into retinal and a di-aldehyde

There are other examples of this secondary cleavage occurring, but this is the most prominent. A similar synthetic compound, β-apo-8’-carotenal (Figure 1-10), can be easily cleaved by CCD enzymes and this is the only example of a synthetic substrate being accepted by a member of the CCD family.38

Figure 1-10. Synthetic substrate β-apo-8’-carotenal

Carotenoid cleavage dioxygenase 1 (EC: 1.14.99.n4)

Carotenoid cleavage dioxygenase 1 (CCD1) utilizes molecular oxygen to cleave carotenoids symmetrically at the 9/10 and 9’/10’ positions to generate di-aldehydes and ketones

(Figure 1-11).38 It is one of the only enzymes in the CCD family to be found in the cytosol.42

29

Figure 1-11. Cleavage of β-carotene at the 9/10 position to give β-ionone and β-apo-10’- carotenal

This enzyme has been the most studied and best characterized in the CCD family. It is also the main focus of our project. The compounds produced by this enzyme are important flavor and fragrance volatiles, e.g., β-ionone. Among the various plant CCD1 enzymes, Arabidopsis thaliana (AtCCD1) is the most studied and well understood, although significant research has also been devoted to Zea mays CCD1 (ZmCCD1) .36, 38 Both of these enzymes were used in this study.

CCD1 can accept a wide range of all-trans and 9-cis-carotenoids, as well as epoxycarotenoids.41 Substrates known to be accepted by this enzyme include lycopene, β- carotene, δ-carotene, zeaxanthin and ζ-carotene (Figure 1-12).43 In a rare occurrence, CCD1 was found to have preferential cleavage of lycopene at the 5/6 and 5’/6’ position.44 This is the only known compound to date that CCD1 does not prefer to cleave at the 9/10 or 9’/10’ alkene bond.

In addition, rice CCD1 (OsCCD1) was found to cleave the 7/8 position of apolycopenals, forming C10-aldehyde geranial (Figure 1-13).44-45

30

Figure 1-12. Carotenoids accepted by CCD1 and their cleavage positions a) lycopene, b) β- carotene, c) zeaxanthin, d) δ-carotene and e) ζ-carotene38

Figure 1-13. Apo-10’-lycopenal being cleaved by OsCCD1 at the 7/8 alkene to form geranial45

Carotenoid cleavage dioxygenases 7 (EC:1.13.11.68) and 8 (EC:1.13.11.691)

Carotenoid cleavage dioxygenase 7 (CCD7) also catalyzes the cleavage of carotenoids at the 9/10 positions.41, 46 The main substrate for this enzyme is β-carotene. 47 While the CCD1 enzyme cleaves symmetrically at the 9/10 and 9’/10’ positions, CCD7 is thought to only cleave asymmetrically.48

31

Carotenoid cleavage dioxygenase 8 (CCD8) cleaves mainly β-apo-10’-carotenal at the

13/14 position, which is the product of β-carotene cleavage from CCD1 and CCD7.41 It is believed that CCD7 and CCD8 act sequentially to produce compounds (such as carlactone) involved in the regulation of shoot branching (Figure 1-14).35, 49 The stereo-configuration

(cis/trans) of the β-apo-10’-carotenal produced by CCD7 ultimately determines the nature of the

CCD8 product and is critical for identifying and understanding the reactions leading up to carlactone synthesis.46 In support of this theory, loss-of-function mutants of either CCD7 or

CCD8 have a developmentally different phenotype.48

Figure 1-14. Sequential cleavage of CCD7 and CCD8 to produce carlactone

32

CCD Enzyme Structure

CCD1 contains an FeII ion that is bound by four highly conserved His residues (Figure 1-

15).16 The iron is octahedrally coordinated to the four His residues and a water; the sixth site is thought to accept molecular oxygen during the catalytic cycle.50 It is assumed that the dioxygen will displace the water molecule once the substrate is bound, coordinating to two of the sites on the iron center. In the crystal structure, the sixth site is unoccupied but this site is not suitable for a water molecule due to the proximity of the methyl group of Thr136.50-51 An iron coordinated to four His side chains is only known in a few other enzymes: mammalian 15-lipoxygenase (PDB

ID: 2P0M), the photo synthetic reaction center of Rhodopseudomonas viridis (PDB ID: 5PRC) and Rhodobacter sphaeroides (PDB ID: 1OGV), and photosystem II of the cyanobacterium

Thermosynechococcus elongates (PDB ID: 1W5C).52

Figure 1-15. The octahedrally coordinated Fe in the active site of the ACO enzyme (PDB ID: 2BIW)

The only member of the CCD1-family with x-ray structural information available is apocarotenoid oxygenase (ACO) from Synechocystis sp. PCC 6803 (PDB ID: 2BIX).50 Although it was soluble, this enzyme could not be crystallized without the aid of a detergent, octylpolyoxyethylene. It was found that the enzyme consisted of a seven-bladed β-propeller with the four His ligands at the axis that holds the FeII ion at the active center (Figure 1-16a). Three of

33

these His residues further hydrogen bond with second-shell glutamates (Glu).51 As can be seen in

Figure 1-16b, there is a deep pocket on the bottom of this enzyme that is thought to be the pathway taken by dioxygen to reach the catalytic Fe; however, this pocket does not extend all the way to the Fe.50 It is believed that other enzymes in this family share a common chain fold, similar active sites, and follow the same reaction mechanism.

(A) (B)

Figure 1-16. Crystal structure of ACO (PDB ID: 2BIW) (A) from the bottom looking up the axis through the enzyme and (B) from the side to see the seven-bladed propeller and loops.

The loops on the top of the propeller (Figure 1-16b) are long and form a large dome over the active center, whereas the loops on the bottom of the structure are very short.50 There is a hydrophobic patch on the surface of ACO that consists mostly of protruding leucines and phenylalanines (Figure 1-17).36 This hydrophobic pocket is thought to dip into the membrane to extract nonpolar substrates, such as carotenoids, making this class of proteins membrane associated.

34

(A) (B)

Figure 1-17. Hydrophobic surface of ACO (A) Cartoon showing the active site tunnel, covered by the dome and a hydrophobic patch41 (b) Hydrophobic patch (red) at the entrance to the active site tunnel. The white residues are the hydrophilic side chains.

As seen in Figure 1-17a, ACO is not able to accept substrates with β-ionone rings on both ends of the molecule. Substrates with a single β-ionone ring, however, are able to enter the active site tunnel for cleavage. This tunnel is also long enough to accommodate apocartenoid substrates. The bond to be cleaved is located on top of the catalytic FeII and then exits through the other end labelled ‘exit’ in Figure 1-17a.41 Kloer et al. propose that the straight isoprenoid tail of carotenoid compounds will enter the tunnel and then collide with the oxygen ligands associated with the FeII center. This likely sterically enforces the conversions from trans to cis and thereby explains how an all-trans substrate can enter the active site tunnel and then become kinked (Figure 1-17a) in order to fit the tunnel for cleavage to occur.50 This isomerization only occurs at methyl substituted double bonds in a conjugated system upon binding.50 After cleavage, the aldehydes have been known to readily convert back to all-trans to exit the tunnel.50

35

A homology model was created for ZmCCD1 by Simon (a previous member of the

McCarty group at the University of Florida) and was based on the crystal structures of ACO and

VP14 (Figure 1-18). As would be expected, this structure closely resembles that of the ACO crystal structure and the same active site features are likely present.

Figure 1-18. Homology model of ZmCCD1

Mechanism

2+ While Fe and O2 are clearly required for carotenoid cleavage, the nature of the reactive oxygen species formed during the catalytic cycle has not been established conclusively.41

Carotenoid compounds are highly reactive due to their extended conjugated double bonds and oxygenation and cleavage can occur by a number of non-enzymatic processes. While it is clear that the scissile C-C bond is cleaved to the corresponding aldehyde and ketone by molecular oxygen, whether the reaction involves a mono- or a dioxygenase pathway has not been established (Figure 1-19).41

36

Figure 1-19. Proposed pathways for carotenoid cleavage by CCD enzymes

Monooxygenases activate molecular oxygen and incorporate one oxygen atom into the substrate while the second is lost as water. The proposed monooxygenase mechanism for CCD involves initial epoxidation of the scissile alkene followed by ring-opening to the diol by water

(18O labelled in Figure 1-19). The diol then undergoes oxidative cleavage by the FeIV oxo species.

Dioxygenases incorporate both oxygen atoms of molecular oxygen into the products.16 In the case of CCD, this would likely involve the formation of a dioxetane intermediate that disproportionates into the two carbonyl species.

In the monooxygenase pathway, one carbonyl product would contain an O2-derived oxygen while the other would contain a solvent-derived oxygen. By contrast, the carbonyl oxygens of both products would contain atoms derived from O2 in the dioxygenase pathway. To

37

determine the actual mechanism, isotopic studies with labelled O2 and H2O have been done.

Unfortunately, rapid exchange of oxygen between the aldehyde carbonyl (via the hydrate) and water solvent complicates the analysis. It is for this reason that this class of enzymes are often referred to as carotenoid cleavage oxygenases (CCOs).

A study done by Leuenberge et al. claimed that β-15-15’-carotenoid cleavage oxygenase

(β-CCO) followed a monooxygenase pathway.53 To prevent the potential oxygen exchange with water, horse liver alcohol dehydrogenase (HLADH) was used to convert the aldehydes to their

17 corresponding alcohols in situ, which were silylated for MS analysis. Using O-labeled O2 and

18O-labeled water, they found that one oxygen atom (17O) was from molecular oxygen and the other from water (18O), thereby supporting the monooxygenase type mechanism. These results were, however, questioned due to the long reaction time (7.5 hours) and the dismutase activity of the HL-ADH, especially at increased levels of NADH, which could lead to unspecific water- derived oxygen incorporation into the products.16

A later study by Schmidt et al. utilized A. thaliana CCD1 (AtCCD1).40 β-Apo-8’- carotenal was used as the substrate because its product, β-ionone (Figure1-8a), should exchange its carbonyl oxygen with water at a very low rate.16, 40 The reaction also required only 30

18 minutes, which is significantly shorter than the prior study (7.5 hours). In the presence of O2,

18 the β-ionone was 96% labeled. In the complimentary experiment with H2O, the β-ionone was unlabeled.40 While there was some oxygen exchange during the reaction (as was shown by a

18 blank reaction), 27% of the C17 dialdehyde showed incorporation of one O atom. These results support a dioxygenase-type mechanism.

While there have been several other attempts to determine the mechanism of action of

CCDs, a firm conclusion has not yet been reached.54 A computational study was carried out

38

using the crystal structure of an AtCCD1 related enzyme, ACO from Synechocystis sp. PCC

6803. Borowski et al. used DFT (B3LYP) to evaluate both the mono- and dioxygenase pathways for this enzyme.51 The energy barrier for epoxide formation was 16.6 kcal/mol, which slightly exceeds that of the dioxetane intermediate (15.9 kcal/mol). This is thought to result from the steric effects of Thr 136 in the catalytic site. These results also showed that subtle changes in the active site of these enzymes could favor one mechanism over the other and opened the possibility that a single CCO enzyme could follow both reaction pathways simultaneously, depending on the substrate.16 Further investigations need to be done to fully elucidate the mechanism of this class of enzymes.

Mutagenesis Studies

There have been a few mutagenesis studies done for AtCCD1, mostly to improve organic solvent stability, protein expression, carotenoid conversion, or to understand structure-function relationships.36, 55 Behrendt, Schwaneberg, and Schrader found that by mutating Ser 363 in

AtCCD1 to Thr, the variant cleaved zeaxanthin with a 1.6-fold increase in conversion as compared to the wild type.36 In addition, when Ser 363 was mutated to Gly, the organic solvent tolerance of AtCCD1 increased by 1.5-fold as compared to the wild type. Ser 363 is predicted to be a surface exposed residue, based on homology models, that is believed to be near the active site entrance tunnel. Finally, it was determined that mutating Ile 10 to Val increased protein expression.36 For this reason, all of our studies used the I10V variant of AtCCD1 to maximize protein yield.

Applications of CCD1

As previously mentioned, carotenoids are the primary substrates for this class of enzymes. This study investigated the possibility that unsaturated fatty acids might also be accepted as substrates for CCD1. Several fatty acids such as linoleic acid, linolenic acid, and

39

arachidonic acid have multiple degrees of unsaturation and have no functionalization, which might allow them to access the active sites of these enzymes. There is one example reported in the literature stating that CCD1 can accept an unnatural substrate, β-apo-8’-carotenal (Figure 1-

20).38, 40, 55 The structure of this particular compound is not symmetrical like typical carotenoids, suggesting that the substrates accepted by CCD1 do not necessarily have to be symmetrical to be cleaved. This observation is further supported by the cleavage of 9-cis-neoxanthin.55 Several substrates that are known to be cleaved by AtCCD1 are shown in Figure 1-20.38, 41, 44, 55

Carotenoids are very hydrophobic, a property shared by long-chain unsaturated fatty acids and also have the characteristic long carbon chains, similar to carotenoids

Figure 1-20. Several substrates known to be cleaved by AtCCD1

When linoleic acid is aligned with two known substrates for CCD1, the C12-C13 alkene appears to be positioned similarly to the alkenes known to be cleaved in normal CCD1 substrates

40

(Figure 1-21). Cleavage of the C12-C13 alkene in linoleic acid would yield n-hexanal in a single- step process.

Figure 1-21. Linoleic acid aligned with two known substrates of CCD1

The possibility of using CCDs for industrial purposes has drawn a lot of attention.16 To date, products derived from CCDs are not made in sufficient quantities to warrant commercial processes. If the yields could be improved, this class of enzymes has the potential to revolutionize the flavor and fragrance industries. Schilling et al. reported an increase in expression of AtCCD1 in E. coli by adding a glutathione-S-transferase (GST) fusion tag to the protein.56 By using this tag in conjunction with Triton X-100 and 15% methanol as organic co- solvent an 18-fold increase in activity was observed. In contrast, Vogel et al. reported that a GST fusion could inhibit CCD activity. While there is still a long way to go before an industrial process can be considered, this observation argues that it may one day be a possibility.

Lipoxygenases

Identification and Characterization

Lipoxygenases (LOX: EC 1.13.11.12) are non-heme, metal dependent dioxygenases.57

While the inorganic cofactor is predominantly iron, there are a few cases where manganese is

41

used, including lipoxygenases from Gaeumannomyces graminis, Magnaporthe salvinii,

Aspergillus fumigatus.58-59 These enzymes are widely distributed in the plant and animal kingdoms. There have been a few examples seen in the microbial kingdom; however, these have received less attention.58 The main difference between plant and animal lipoxygenases is their positional selectivities. Plants generally produce 9- and 13-LOXs, which mainly use linoleic acid as a substrate.60 Animal LOXs generally are 5-,8-,12-, and 15-LOXs and typically utilize arachidonic acid as a substrate.61 LOX enzymes are multifunctional, catalyzing at least 3 distinct reactions: (a) deoxygenation of lipid substrates, (b) conversion to hydroperoxy lipids, and (c) formation of epoxy leukotrienes.34, 57 In general, they catalyze the oxidation of unsaturated fatty acids at the corresponding carbon that contains a cis, cis-1,4-pentadiene system such as the moieties found in linoleic (18:2), linolenic (18:3), and arachidonic acids (20:4) (Figure 1-22).7a

The final product is a peroxide containing fatty acid (Figure 1-5), which can be reduced enzymatically by glutathione peroxidase or by a chemical reductant such as Tris(2- carboxyethyl)phosphine (TCEP).62

Figure 1-22. Cis, cis-1,4-pentadiene systems (a) linoleic acid, (b) linolenic acid, and (c) arachidonic acid

LOX enzymes have significant physiological functions and are receiving attention for their roles in asthma and allergies.63 The most abundant source of lipoxygenase can be isolated

42

from soybean seeds.64 Axelrod et al. developed a procedure for isolating soybean lipoxygenase that is commonly employed.64 They also found that this source contains three isozymes: LOX1,

LOX2 and LOX3 that can be differentiated on the basis of pH optima, substrate specificity, and product formation.7a Each of the soybean LOX isozymes is listed in Table 1-3 along with their

64 pH optima, KM values, and regioselectivities.

Table 1-3. Key differences between the isozymes of soybean LOX enzymes64

Enzyme pH optimum KM (mM) Regioselectivity LOX1 9 to 9.5 0.012 13-hydroperoxide* LOX2 6 to 7 0.0096 (pH 6.8) 9 and 13 hydroperoxides in equal amounts LOX3 6 to 7 0.34** 65% 9-hydroperoxide 35% 13-hydroperoxide *When Arachidonic acid is used as a substrate, the 15-hydroperoxide is formed first **Apparent KM value due to significant substrate inhibition

It is worth mentioning that there have been substrates known to be accepted by LOX enzymes besides free unsaturated fatty acids that include acylglycerols, phosphoglycerides, phospholipids, cholesterol esters, allylic ketones, glycerides, alcohol sulfates, hydroxamates, polyunsaturated alkyl-1-halides, and even soybean biomembranes.33-34, 64-65 While these are considered substrates for LOX, their oxidation rates are much lower than those of free fatty acid.33 Reactions with acylglycerols, phosphoglycerides, and phospholipids also need to have a bile salt present such as deoxycholate due to their low solubility. The mechanism of how these more complicated substrates bind in the active site needs elucidation and more structural determinations may be required.34

Enzyme Structure

In 1996, Minor et al. reported the crystal structure of LOX-1 at 100K and a 1.4Å resolution (Figure 1-23).66 Plant LOX enzymes are 95-100 kDa and consist of two domains. The

43

amino-terminal end (residues 1-146) is a β-barrel domain that is between 25-30 kDa and structurally similar to C2 domains.57 This β-barrel is thought to be involved in membrane binding and is not necessary for catalytic function; however, when deleted, the activity of the mutant is lower than the wildtype protein.67 The carboxy-terminal end (residues 147-839) is about 55-65 kDa and consists mostly of α-helices that harbors the catalytically active iron and the active site.68 In soybean LOX1, helix 9 (residues 473-518) is the central structural element and is 65Å long.66 In Figure 1-23, this helix is shown in dark green (indicated by arrow) down the center of the structure.

Figure 1-23. 1.4Å crystal structure of LOX-1 (PDB ID:1YGE)

The catalytic iron is octahedrally coordinated by 5 amino acid residues in the active site and a hydroxide ligand (Figure 1-24).57, 68 This hydroxide ligand is thought to come from a water molecule present in the active site. The carbonyl oxygen of Asn 694 is 3Å from the iron, implying a relatively weak coordination interaction; however, a hydrogen bonding network

44

coordinates the carbonyl and iron which can be observed in the crystal structure (Figure 1-

24b).66, 69 While Asn 694 lies in the first coordination sphere of the iron, the second coordination sphere is made up of Gln 495 and Gln 697.69 The crystal structure determined by Minor et al. showed the resting state of the enzyme, with the catalytic metal in the Fe2+ state due to the low temperature used in data collection (100K).66

(a) (b) Figure 1-24. Octahedrally coordinated iron in the soybean LOX1 active site (a) showing coordination to each of the ligands and (b) adapted from the 1.4Å crystal structure66

Several studies have been conducted to determine the residues that determine the regiospecificity of LOX enzymes. In the substrate orientation model, it is theorized that lipids penetrate the active site with their methyl end first (straight orientation) in 13-LOX and carboxyl end first in 9-LOX (inverse-fashion).57 Minor et al. determined that an Arg residue present in the substrate-binding pocket may form a salt bridge with the carboxyl group of the lipids when bound in the inverse fashion.66 Plant LOX enzymes, however, have a highly conserved Phe or

His residue that shields the positive charge of the Arg from the carboxyl group (Figure 1-25a).34

It is thought that the Phe or His residues prevent the carboxyl end of the lipid from penetrating the active site since there is no counterpart to neutralize the negative charge. Feussner and Kühn found that Phe or His is highly conserved in plant 13-LOX enzymes. In the 9-LOX enzymes, however, this residue is nearly always Val (Figure 1-25b).70 When they mutated the His residue

45

in cucumber 13-LOX to Val and Met residues, the regioselectivity of the mutants were altered.71

Finally, there is a highly conserved lysine (Lys) residue in plant 13-LOX enzymes that is thought to form a salt bridge to the carboxyl group of the lipids when bound in the straight orientation.34

Figure 1-25. Active site depiction of soybean LOX-3 harboring linoleic acid34, 72 (A) carboxyl group of linoleic acid (LA) near the Arg residue that is shielded by Phe643/834. (B) Phe residue (green) of 13-LOX and Val residue (blue) of 9-LOX determining regiospecificity of the free fatty acids.

A second model worth mentioning is called the space related model based on data from a mammalian 15-LOX enzyme.73 In general, the fatty acid penetrates the active site with its methyl end first and the depth of the substrate binding pocket determines the site of hydrogen abstraction and the where the molecular oxygen will insert.34, 57 In plant LOX enzymes, however, it seems that only one double allylic methylene group from free fatty acids is accessible to the enzyme thus rendering this model unlikely.

Mechanism

As mentioned previously, LOX enzymes are multifunctional, catalyzing at least three distinct reactions.34, 57 The first step in the oxygenation mechanism is hydrogen abstraction. This

46

is quickly followed by a radical rearrangement (resonance structure) and finally oxygen insertion

(Figure 1-26). 34, 57, 74 The mechanism shown below is for 13-LOX, but the 9-LOX enzyme follows an analogous pathway at the 9 carbon. The final product of this reaction is either (13S)- hydroperoxy (Figure 1-26) or the (9S)-hydroperoxy derivative of the free fatty acid, in this case linoleic acid.

Figure 1-26. The 13-LOX enzyme mechanism and its regioselectivity using linoleic acid34, 57

It is thought that catalysis is initiated by reaction of the iron (III) form of the enzyme with the fatty acid substrate, which then reacts with dioxygen.74 There are three proposed pathways in which this radical-mediated reaction can occur (Figure 1-27).74 Clapp et al. determined through radical trapping experiments that pathway A is the more likely route of radical rearrangement. A monounsaturated fatty acid was oxygenated 4 to 5 orders of magnitude more slowly than linoleic acid. The absence of either double bond forces the rearrangement to occur through an allylic radical rather than the more stable pentadienyl radical.74 In pathways B and C, one of the two double bonds is more critical than the other. The experimental evidence thus far shows that the two double bonds contribute equally to the C-H abstraction, therefore pathway A is more likely.

47

In addition, a density functional theory calculation suggested that the pentadienyl radical in pathway A is the more favorable mechanism.75

Figure 1-27. The three proposed radical rearrangement mechanisms for the 13-LOX enzyme74

Lineolate 13-Hydratase

Identification and Characterization

Hydrated fatty acids are naturally found as components of triglycerols, cerebrosides, and other lipids in plants, animals, insects, and microorganisms.76 They are commonly used as starting materials for the synthesis of resins, plastics, nylons, and biopolymers.77 Biopolymers made from these compounds have advantages over petroleum-based polymers due to their higher resistance to heat, chemicals, and impact. They also have increased flexibility, higher biocompatibility, and no toxicity.77 In addition, they can be used as additives for emulsifiers, lubricants, stabilizers, coatings, and paint.76 Finally they can be used for precursors of lactone production. It is widely known that hydroxylated fatty acids have increased reactivity, viscosity,

48

and solubility as compared to their non-hydroxylated counterparts. They have been found to have antibiotic, anti-inflammatory, and anticancer activities as well.77-78

In 1992, Koritala and Bagby determined that linoleic acid could be hydrated at the C10 position using whole cells of Nocardia cholesterolicum (NRRL 5767) (Figure 1-28).79 When hydrating oleic acid, full conversion was reached in 8 hours and only 70 mg of cells (dry weight) were required. With linoleic acid, the conversion tapered off after 8 hours at 71% and by 24 hours had only reached 77%. This reaction also required roughly 350 mg of cells (dry weight). It is thought that the lower conversion was due to product inhibition. They also determined that the optimum pH and temperature for this reaction was 6.5 and 35°C, respectively. In 1998, Hudson et al. determined that linoleic acid could be hydrated at both the C10 and C13 position using cells of a ruminant bacterium, Streptococcus bovis (JB1) (Figure 1-28).80 This was the first report of a position 13-selective hydration. Kishimoto et al. reported in 2003 that L. acidophilus IFO 13951 cells regiospecifically made 13S-hydroxy-cis-9-octadecenoic acid from linoleic acid.81

Figure 1-28. Hydration of linoleic acid to form the 13- and 10- hydroxyl products

Since these initial studies, several other organisms were found to hydrate linoleic acid.81

Lactobacillus plantarum, Streptococcus bovis, and Flavobacterium sp. DS5 (NRRL B-14859) were among those found to produce 10-hydroxy-12Z-octadecenoic acid from linoleic acid.80, 82

Lactobacillus acidophilus, Pedicococus pentosaceus, and P. sp. AKU 1080 were all found to

49

produce 13-hydroxy-9Z-octadecenoic acid.81, 83 In most cases, both isomers were produced from one organism.84

In 2015, Hirata et al. screened various lactic acid bacteria (more than 300 strains) for the ability to convert linoleic acid into hydroxy-linoleic acid.85 A majority of these strains converted linoleic acid into 10-hydroxy-12Z-octadecenoic acid and some produced conjugated linoleic acid isomers. It was determined that L. acidophilus NTV001 has the highest ability to complete the conversion to 13-hydroxy linoleic acid (96%). The gene responsible for this conversion was named fatty acid hydratase 1 (FA-HY1). Interestingly FA-HY1 produced 13R-hydroxy-9Z- octadecenoic acid, rather than the (S)-configuration observed for L. acidophilus IFO 13951.81

FA-HY1 was found to have a broad substrate range and catalyzed the hydration of C16, C18, C20 and C22 fatty acids. This was the first reported conversion of C20 and C22 fatty acids by a bacterial enzyme. A second hydratase was found to be associated with L. acidophilus NTV001, which was named FA-HY2. It was found to produce only 10S-hydroxy-12Z-octadecenoic acid.

The observation that two hydratase enzymes are associated with a single organism has also been made previously.82b, 84

In 2015, Oh et al. found that L. acidophilus NBRC 13951 selectively produced 13- hydroxy linoleic acid.84 This was the first report of a lineolate 13-hydratase being cloned to produce the hydroxylated product. The gene was inserted into plasmid pET-15b and used to transform E. coli ER2566 to yield the final overexpression construct. Purified enzyme (1.5 mg/mL) was used at a pH of 7.0 and 35°C to transform linoleic acid into 13-hydroxy-9- octadecaenoic acid. The product was made at a titer of 8 mM after 3 hours of reaction time. To make 10-hydroxy-12Z-octadecenoic acid, a different gene was used along with 2.0 mg/mL of

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purified hydratase. In this study, the hydroxy-linoleic acid compounds were further used to produce lactone products using whole cell biocatalysis and wild type yeast cells.

Park et al. conducted a second study to optimize the reaction conditions for the lineolate

13-hydratase assay.77 This was done utilizing the same expression system as Oh et al. The optimal conditions were determined to be pH 6.0, 40°C, 0.25% (v/v) Tween 40, 25 g/L cells, and

100 g/L linoleic acid. Under these conditions, 79 g/L of 13-hydroxy-9Z-octadecenoic acid was produced in 3 hours, which represented a 79% (w/w) conversion. This is 35- and 59- fold higher than the amount produced under non-optimized conditions.84 The regioselectivity was also determined to be 96% 13S-hydroxy linoleic acid, matching that determined previously.81

Enzyme Structure

Lineolate 13-hydratase has only been discovered recently and relatively little structural information is available. In 2013, Volkov et al. determined the crystal structure of a fatty acid double bond hydratase from L. acidophilus NCFM (LAH) to a resolution of 2.3Å without a cofactor (Figure 1-29a) and 1.8Å with bound linoleic acid (Figure1-29b).86 This hydratase is known as a myosin cross-reactive antigen (MCRA), which forms a large family that is broadly conserved across various bacteria.

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Figure 1-29. Crystal structure of a fatty acid double bond hydratase from L. acidophilus (A) with no cofactor (PDB ID: 4IA5) and (B) with linoleic acid (green) bound (PDB ID: 4IA6)

The LAH structure is a homodimer and each protomer consists of four domains.86 Three of these domains (residues 1-538) form the FAD-binding and substrate binding sites. The fourth domain (residues 539-591) is located at the C-terminus and consists of three α-helices connected by two loops. It covers the hydrophobic substrate cleft leading to the active site. The fourth domain undergoes conformational changes once linoleic acid is present, opening the entrance to the active site cleft (Figure 1-30a). This cleft contains the FAD-binding domain. These helices block the entrance to the active site otherwise (Figure 1-30b). As seen in Figure 1-30, the residues on the surface of the protein are mostly hydrophilic, suggesting that this is not a membrane associated protein. It is thought that the entrance to the hydrophobicity of the active site cleft attracts the non-polar substrates.

Figure 1-30. Hydrophobicity plot of an LAH (A) Entrance of the hydrophobic tunnel (red) to the active site. Linoleic acid is shown in cyan entering the tunnel. The white residues shown are hydrophilic. (B) The entrance to the tunnel is blocked.

Oleate hydratase is the second enzyme in this family which structural data is available.

Engleder et al. solved the crystal structure of oleate hydratase from Elizabethkingia meningoseptica in the presence of FAD to 2.75Å resolution (Figure 1-31). This enzyme shows

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41.6% sequence similarity with lineolate 13-hydratase from L. acidophilus KCTC 3164 (the enzyme used in our study).87 This sequence similarity is sufficiently high to draw inferences about the core structure of the latter.

Figure 1-31. Crystal structure of Oleate hydratase from Elizabethkindia meningoseptica (PDB ID: 4UIR) with the FAD (green) on the left side of the protein

Oleate hydratase crystallized as a homodimer, similar to the double bond hydratase mentioned previously with the FAD cofactor in chain A (orange in Figure 1-31).87 Poor electron density was shown in chain B for FAD (blue in Figure 1-31), indicating a low occupancy. A major difference in cofactor binding was observed for a loop region in the binding pocket. In the presence of the cofactor, it is well ordered; however, in its absence, less order was observed, suggesting a structural role for cofactor binding. A hydrophobic cavity was also observed close to the cofactor, similar to the double bond hydratase. The cavity adopts a V shape, with one end being more hydrophilic due to the presence of residues Gln 265, Thr 436, Asn 438, and His 442

(Figure 1-32).

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Figure 1-32. Active site of Oleate hydratase with the FAD cofactor in yellow and the oleic acid in cyan87

The cis double bond of oleic acid is located at the bend in this cavity, with the carboxylate close to the previously mentioned hydrophilic residues. Residues Glu 122, Tyr 241, and Tyr 456 are located in close proximity to the double bond that is hydrated.

Mechanism

There have been few mechanistic studies on this class of enzyme. Engleder et al. performed mutagenesis studies on the various active site residues (Figure 1-32) to propose a mechanism for oleate hydratase.87 Based on this data, Tyr 241 is thought to protonate the double bond and Glu 122 is thought to activate a water molecule for attack on the partially charged double bond (Figure 1-33). This is consistent with complete loss of catalytic activity when either residue was mutated to Ala. Glutamate 122, is located on a flexible loop that becomes ordered upon cofactor binding.

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Figure 1-33. The proposed mechanism for alkene hydration adapted from Engelder et al.87

As mentioned previously, the FAD cofactor is thought to play a structural role for this enzyme. Based on isothermal titration calorimetry (ITC) studies, it was determined that FAD

87 was non-covalently bound to the enzyme with a KD value of 1.8 μM. It is also believed that reduced FAD is present in catalysis. Loss of FAD resulted in a loss of enzymatic activity.

Activity was restored to 80% by re-adding FAD. Several roles have been proposed for this cofactor including enzyme stabilization, correct localization of the substrate, positioning the amino acids for catalysis, or direct participation in the mechanism. Studies to confirm the proposed mechanism and determine the apparent role of FAD remain to be carried out.

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CHAPTER 2 USING CAROTENOID CLEAVAGE DIOXYGENASES TO CLEAVE LINOLEIC ACID INTO N-HEXANAL

Background

Our initial attempt at naturally producing n-hexanal used the carotenoid cleavage dioxygenase 1 enzyme, which is predicted to cleave the 12/13 alkene bond in linoleic acid

(Figure 2-1). This route was attractive due to it having a single step and only using molecular oxygen as a cofactor.

Figure 2-1. Predicted linoleic acid cleavage position of CCD1 into n-hexanal

There were several problems that needed to be solved upon starting work with CCD1 including n-hexanal and linoleic acid quantitation, enzyme preparation, optimal reaction conditions, autoxidation prevention and enzyme activity determination. The following chapter explores each of these problems and the associated solutions that were developed in for each instance.

Experimental Strategy n-Hexanal Quantitation

Our initial task in this project was to quantitate the n-hexanal synthesized from reactions catalyzed by CCDs. The high reactivity, partial water solubility and significant vapor pressure of n-hexanal conspire to make this a challenging analyte. We first explored solid phase micro extraction (SPME), which has been reported as the preferred method for n-hexanal quantitation and for volatiles in general.88 While we successfully detected n-hexanal using SPME, the very low throughput was a serious drawback and its reproducible quantitation proved impossible to

56

achieve in our hands. We then attempted to quantitate n-hexanal using DNPH derivatization.

DNPH derivitization is used to derivitize aldehyde compounds and make them visible on high pressure liquid chromatography (HPLC) with UV detection.89 While the derivatization procedure worked on pure n-hexanal, it was determined that there was not enough produced from CCD biotranformations for accurate quantitation using DNPH. We therefore turned to solvent extraction as a means of removing n-hexanal. These efforts included biphasic reactions (aqueous buffer mixed with an immiscible organic solvent) as well as post-reaction solvent extraction. The latter proved most successful when paired with an internal standard. Several were evaluated and methyl benzoate was found to be most suitable for n-hexanal quantitation using GC/MS analysis.

CCD Production and Isolation

We evaluated two cloned CCDs: Zea Mays CCD1 (ZmCCD1) and Arabidopsis thaliana

CCD1 (AtCCD1). The former was received as a generous gift from the McCarty lab (University of Florida) in a pGEX-2T vector. Unfortunately, the expression level of active ZmCCD1 was relatively low and we had difficulty trying to enhance its activity. We therefore turned to the better-characterized AtCCD1 gene, which was synthesized chemically and cloned into pUC57 plasmid. Two different affinity tags were appended to the AtCCD1 gene for purification purposes: glutathione S-transferase (GST) and maltose binding protein (MBP). The GST tag interfered with AtCCD1 oxidations of linoleic acid and it could not be easily removed. We suspected that n-hexanal might be consumed by reactions with exposed amino groups on the protein, so both acetylation and cross linked enzyme aggregates (CLEA) were explored. The

MBP tag proved more efficient for AtCCD1. This tag could be easily cleaved from the AtCCD1 fusion protein and it also increased the solubility of the overexpressed protein. In a final attempt to increase the overexpression of AtCCD1, the plasmid was transformed into the BL21 (DE3)-

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pLysS strain, which is preferred for proteins that are somewhat toxic to E. coli. The purity and expression levels were evaluated using polyacrylamide gel electrophoresis.

Cleaving Linoleic Acid to n-Hexanal Using CCD1 Enzymes

Initial studies using AtCCD1 to convert linoleic acid into n-hexanal used the procedure of

Vogel et al.38 Several variations of this procedure were performed in order to optimize our

AtCCD1 reactions.

Experimental Procedures

Linoleic acid solution

A 1% Tween-20 solution was made by adding 10 µL of Tween-20 (Fisher BP337-500) to

1 mL of Milli Q water. A 30 mM linoleic acid solution was then prepared by adding 9.35 µL of linoleic acid (Acros 60-33-3) to the 1 mL 1% Tween-20 solution. The solution was sonicated for

1 minute and became cloudy white when the linoleic acid had been sufficiently dispersed.

Light Scattering Experiments

These were done in collaboration with the Savin group at the University of Florida. A 30 mM stock of linoleic acid was prepared as previously described. Various dilutions of linoleic acid were prepared ranging from 0.001-0.05% making sure that the samples remained non viscous. Each light scattering cell was cleaned multiple times to remove any dust particles that may have been present. The dilutions were filtered into individual sample cells and placed into the instrument to collect light scattering data from various angles. The same procedure was followed for various dilutions of AtCCD1 ranging from 0.01-0.05%. Finally the same AtCCD1 dilutions were prepared with 0.1% Triton X-100 added.

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n-Hexanal Quantitation

1. Solid phase micro extraction (SPME)

The SPME procedure was done manually. The holder (for sampling) and the fibers were purchased through SUPELCO (Bellefonte, PA, USA). Three fibers were evaluated with different coatings: PDMS/DVB with 65 μM thickness, CAR/PDMS with 75 μM thickness, and

DVB/CAR/PDMS with 50/30 μM thickness. They were conditioned prior to use according to the manufacturer instructions (Table 2-1). The individual parameters were set on the GC followed by a 1 hour baking time period.

Table 2-1. Conditioning parameters for each of the SPME fibers evaluated Inlet Detector Oven Fiber temperature temperature temperature Color PDMS/DVB 250˚C 280˚C 280˚C blue CAR/PDMS 300˚C 300˚C 300˚C black DVB/CAR/PDMS 270˚C 280˚C 280˚C grey

Aqueous 1 mL samples to be analyzed were placed into a crimp top GC vial with a small magnetic stir bar and 1 mM methyl benzoate (internal standard). The vial was capped and put into a heat block set at 75°C using the SPME holder for 30 minutes (equilibration time). The fiber was inserted into the vial and exposed to the head space for 30 minutes (extraction time).

The fibers were immediately inserted into the injector port on the GC/MS for thermal desorption for 5 minutes prior to analysis using EDA2_MTH. To ensure that all impurities were removed from the fibers, blank runs were performed between each sample analysis and each run was completed in triplicate. Care was taken to avoid fiber contact with the aqueous solutions.

GC/MS analysis using EDA2_MTH

Gas chromatography-mass spectrometry was performed on a Hewlett-Packard (HP) 5890

Series II Plus GC (He carrier gas; 1.0 mL/min; split/splitless injector (Table 2-1); injection

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volume 1µL) with a DB-17 (30m long; 250 µm i.d; 0.25 thickness) column. A 0.75 mm i.d. narrow-bore glass liner (Suppelco) was used specifically for SPME fibers to prevent peak broadening and improve the peak resolution. The injection temperature was set to 200°C. The temperature was initially set to 60°C for 5 minutes. The temperature was programmed from 60ºC at 10ºC/minute to 250ºC with a hold for 10 minutes. Manual injections were performed. The GC was coupled to a HP 5971 series mass selective detector. The retention time of n-hexanal

(Aldrich 66-25-1) was determined using standard solutions and verified by comparing its mass spectrum with the NIST library.

2. 2,4-Dinitrophenylhydrazine (DNPH) derivitization

Figure 2-2. DNPH derivitization of hexanal

To prepare a standard, n-hexanal (2 mL) was dissolved in 2 mL of ethanol in a beaker. To this mixture, 2 mL of DNPH reagent (Sigma) was added, stirred, and allowed to stand for 5 minutes. The solid was collected by suction filtration and then was washed with two 5 mL portions of sodium bicarbonate followed by 10 mL of water. The solid was collected in a beaker and 20 mL of 95% ethanol was added, heated until the solid was fully dissolved, and then cooled at room temperature for 5 minutes followed by 10 minutes on ice. The solid was collected by suction filtration and washed with 10 mL of cold ethanol. The synthesized n-hexanal 2,4- dinitrophenylhydrazone was an orange solid (0.5 g, 90%). m.p. 100-102°C, lit. m.p. 101-103°C.90

To further confirm that n-hexanal was sufficiently derivitized, an H1 NMR was taken and its

1 identity confirmed (Appendix D). H NMR (300 MHz, CDCl3): δ 11.01 (s, 1H), 9.12 (s, 1H), 8.3

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(dd, J= 9.7 Hz, 1H), 7.93 (d, J= 9.7 Hz, 1H), 2.43 (m, 2H), 1.62 (m, 2H), 1.38 (m, 4H), 0.93 (s,

3H) ppm.

3. Solvent extraction using d12-n-hexanal

d12-n-Hexanal was added as an internal standard (1 mM) to enzyme reactions prior to solvent extraction (1 mL). The mixtures were then extracted with two 0.5 mL portions of EtOAc.

In addition to d12-n-hexanal, lauric acid and methyl benzoate were also used as internal standards, depending on the reaction system.

Resin recoveries

Several resins were evaluated for n-hexanal recovery. These included XAD4, XAD7 and

(aminomethyl) polystyrene. With XAD4 and XAD7, a small amount of resin was added to an aqueous solution of 5 mM purified n-hexanal. These were rotated at 25°C overnight (>14 hours), the resin was filtered off and then the absorbed material was eluted by washing with methylene chloride. The extracts were analyzed by GC/MS using JON_METH. The (aminomethyl) polystyrene resin was added to an aqueous solution of 5 mM n-hexanal. Samples were rotated overnight (>14 hours) at 30°C and then 1 mM internal standard was added. The solutions were acidified with 3 drops of 1 M HCl before being extracted with two 0.5 mL portions of EtOAc.

The combined extracts were analyzed by GC/MS using JON_METH.

Linoleic Acid Quantitation

FAME derivitization

This procedure was based on that of Ichihara and Fukubayashi.91 Commercial concentrated HCl (37%, w/w, 9.7 mL) was diluted with 41.5 mL of methanol resulting in an 8 %

(w/v) HCl solution. This contains 85% methanol and 15% (v/v) water derived from the concentrated HCl. This solution was stored at 4°C when not in use.

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A 100 μL aliquot of each reaction was placed in a 10 mL glass vial. To the sample, 1.50 mL methanol and 0.3 mL of the 8% (w/v) HCl solution were added in that order. The tubes were vortexed and placed at 45°C overnight (>14 hours) for esterification. Alternatively, the reaction could be performed at 100°C for 1 hour. After cooling to room temperature, 1 mL of hexanes was added followed by 1 mL of water. The tubes were vortexed, the hexanes layer removed, and immediately analyzed by GC using JON_METH.

MSTFA derivitization

Samples were evaporated to dryness prior to analysis on the speed vacuum. Distilled pyridine (2µL) was added to the dried sample along with 50 μL MSTFA (Sigma) and mixed thoroughly. The reactions were incubated at 37°C for 30 minutes with shaking at 250 rpm, then cooled to room temperature and placed into an insert within a GC autosampler vial for GC analysis using EA1_METH. The samples were analyzed immediately since the silyl derivatives degraded rapidly.

GC/MS analysis: EA1_METH

Gas chromatography-mass spectrometry was performed on a Hewlett-Packard (HP) 5890

Series II Plus gas chromatograph (He carrier gas; 1.0 mL/min; split/splitless injector 220ºC; injection volume 1µL) with a DB-17 column (30m long; 250 µm i.d; 0.25 µm thickness). The initial temperature was set to 60°C for 2 minutes (3.5 minute solvent delay). The temperature was programmed from 60ºC at 10ºC/minute to 250ºC (hold for 10 minutes). The GC was coupled to a HP 5971 series mass selective detector. The retention time of linoleic acid (ACROS

60-33-3) was determined using standard solutions and verified by comparing the mass spectrum with that from the NIST library.

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ZmCCD1 Expression

A plasmid containing the ZmCCD1 gene in a pGEX2T plasmid vector was obtained from the McCarty group (University of Florida). The plasmid was used to transform E. coli

BL21(DE3)-Gold cells.38 Individual colonies were grown in 10 mL of LB containing 0.1 mg/mL ampicillin overnight at 37°C. The following day, 10 mL of overnight culture was used to inoculate 1 L LB containing 0.1 mg/mL ampicillin in a baffled flask. The culture was grown

(250 rpm at 37°C) to an OD600 of 0.6 and then induced with 0.1 mM isopropyl β-D-1- thiogalactopyranoside (IPTG). After induction, the culture was grown overnight (250 rpm at

30°C) and centrifuged at 6,000 rpm for 15 minutes. The cells were frozen at -20°C. Frozen cells were thawed on ice and then resuspended in an equal volume of 50 mM NaPO4, pH 7.5. The cells were lysed using a high pressure homogenizer (French Press). Crude lysate samples were stored in aliquots at -20°C in 20% glycerol.

Construction of pEA1

The AtCCD1 gene (Accession number: NM_116217) was synthesized by GenScript and sent to us in a pUC57 plasmid vector. Plasmid pBS2, which contains a GST tag and a T7 promoter, was created by a previous group member (Brad Sullivan).92 Both plasmids were prepared on large scales and purified by CsCl density gradient ultracentrifugation in the presence of ethidium bromide. The AtCCD1 gene was cloned into the pBS2 plasmid vector using NdeI and XhoI restriction enzymes. All linearized products were purified by agarose gel electrophoresis prior to ligation using a 3:1 (vector:insert) ratio. After ligation, the mixture was used to transform E. coli E10Blue cells. Plasmid DNA was isolated from randomly-chosen colonies and sequenced to verify the desired construct, which was designated pEA1. Plasmid pEA1 was used to transform E. coli BL21(DE3)-Gold and BL21(DE3) pLyS strains.

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Construction of pEA2

The pMAL-c5x plasmid vector was obtained from New England Biolabs and purified on a large scale as described above. The previously-isolated AtCCD1 gene was ligated between the

NdeI and EcoRI sites of pMAL-c5x as in the previous example. The resulting plasmid was designated pEA2.

Construction of pEA3

The pET-22b(+) plasmid vector was purified by Dr. Jon Stewart via CsCl density gradient ultracentrifugation in the presence of ethidium bromide. The previously-isolated

AtCCD1 gene was inserted between the NdeI and XhoI sites of pET-22b(+) as described above.

The resulting plasmid was designated pEA3.

AtCCD1 Expression

The appropriate plasmid was used to transform E. coli into BL21(DE3)-Gold strain.

Colonies of the plasmid were grown on LB plates containing 0.2 mg/mL ampicillin and then a single colony was used to inoculate an overnight culture grown at 37ºC (10 mL of LB containing

0.1 mg/mL ampicillin). This was added to 1 L of sterile LB containing of 0.1 mg/mL ampicillin.

This culture was grown at 37°C and 250 rpm until it reached an OD600 of approximately 0.6, then was induced by adding IPTG to a final concentration of 0.1 mM. Once induced, the culture was shaken at 18°C for 48 hours at 250 rpm. The cells were collected by centrifugation at 6,000 rpm,

4°C for 15 minutes. They were resuspended in an equal amount of 50 mM NaPO4, pH 7.5 and lysed using a high pressure homogenizer (French Press). To the lysed cells, 0.2% Triton X-100 was added, mixed thoroughly, and the mixture was centrifuged at 18,000 rpm, 4°C for 60 minutes.56

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Acetylation of AtCCD1

The procedure for acetylating AtCCD1 was based on a method developed by a previous group member (Adam Walton).93 Four microcentrifuge tubes containing 100 μL of purified protein diluted in 400 μL of buffer (0.01% Triton X-100 in 20 mM Tris, 0.2 M NaCl, pH 7.4) were prepared (1-4). Sodium acetate (190 mg) was added to tubes 2 and 4 and neat acetic anhydride (1 μL) was added to tubes 3 and 4. All tubes were rotated at 4°C for 1 hour and then tubes 2-4 were dialyzed against 100 mM KPi pH 7.5, 50 mM NaCl, 50% glycerol for 3 hours.

Aliquots (250 μL) from samples 1-4 were put into microcentrifuge tubes and stored at -20°C until analysis. Each sample was analyzed by a 10% resolution polyacrylamide gel and then taken to the mass spectrometry facility for trypsin digest analysis. The activity of each acetylated

AtCCD1 sample was evaluated using the β-apo-8’-carotenal cleavage assay.

CLEA Preparation

The procedures to make CLEA preparations of AtCCD1 were adapted from Kartal and

94 Kilinc. To determine the optimum (NH4)2SO4 saturation levels, pure AtCCD1 (50 μg) and crude lysate containing AtCCD1 (75 μg total protein) were used. These were brought to specific

(NH4)2SO4 saturations levels between 20 and 80%. These were put on ice for 30 minutes followed by centrifugation for 5 minutes at 15,000 rpm, 4°C. The pellets were weighed after centrifugation and the one with the highest mass is the optimum (NH4)2SO4 saturation level.

Next, the optimum glutaraldehyde concentration was determined. Glutaraldehyde (25% aqueous solution) was added to the 60% saturated (NH4)2SO4 suspensions in concentrations between 12.5 and 250 mM. The vials were rotated at 4°C for 1 hour and then centrifuged at

15,000 rpm, 4°C, 10 minutes. The resulting pellets were resuspended in 100 μL of 50 mM

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NaPO4, pH 7.5. To determine enzyme activity, β-apo-8’-carotenal cleavage assays were carried out.

β-Apo-8’-Carotenal Oxidations

The procedure was adapted from Vogel et al.38 A 0.1 mg/mL stock of β-apo-8’-carotenal

(Sigma) was prepared by dissolving 0.01 g in 100 mL 95% ethanol. This solution was stored in the dark at -20°C under argon when not in use. For enzyme assays, 1 mL of 1% polyoxyethylene sorbitan monopalmitate 20 (Tween 20, Sigma) in 95% ethanol was added to 3 mL of 0.1 mg/mL

β-apo-8’-carotenal solution. The mixture was vortexed vigorously and dried using the vacuum line. Finally, 1.2 mL of 50 mM NaPO4 pH 7.5 was added to the dried residue to make a 250

μg/mL solution.

Assays were performed in a total volume of 1 mL containing 50 mM NaPO4 pH 7.5,

300mM NaCl, 5 μM FeSO4, 5 mM ascorbate, 20% methanol, and 30 μg of β-apo-8’-carotenal.

Water was added to bring the final volume to 0.9 mL and the mixtures were allowed to equilibrate to room temperature. Finally, 100 μL of crude lysate or purified protein was added.

These reactions were mixed, placed on a rotisserie at 25°C in the dark, and allowed to react for 1 hour. The reactions were extracted with 0.5 mL EtOAc and analyzed on HPLC under

EA_MTH1.

For inhibition assays, various substances were added to the AtCCD1 cleavage reaction at a final concentration of 1 mM. These included: linoleic acid, conjugated linoleic acid isomers, methyl lineolate, linolenic acid, arachidonic acid, eladic acid, palmitoleic acid, oleic acid, erucic acid, palmitic acid, lauric acid, hexanoic acid, valeric acid, n-hexanal, n-heptanal, E,E-2,4- decadienal, E-4-decenal, 1-octene, trans crotonic acid, and a triglyceride. This assay was also carried out with β-carotene as a substrate using the same procedure as with β-apo-8’-carotenal.

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HPLC analysis: EA_MTH1

HPLC analysis was performed on a Gilson system consisting of two 306 pumps (A and

B), an 811B dynamic mixer, a PerkinElmer Series 200 autosampler, and a UV/Vis-155 dectector.

A Synergi 4μ Hydro-RP 80A column (4.6 x 75 mm, C18) was used. The following procedure was adapted from Carmen et al.95 The mobile phases consisted of A: acetonitrile/water (80:20) and

B: methyl tert-butyl ether/acetonitrile/water (75:21:4). The gradient elution at 1 mL/min used was 100% A, followed by a linear gradient to 21% B at 7 minutes, a linear gradient to 40% B at

26 minutes, a linear gradient to 80% B at 40 minutes, and finally 80% B isocratically for 4 minutes. A conditioning phase (5 minutes) was then used to return the column to the initial concentrations of A and B. Fifty microliters of each sample were injected and the absorbances at both 441 and 461 nm were monitored.

Protein Purification

Gluathione S-transferase affinity purification

The crude lysate was added to the GST column (1 cm i.d.; 17 cm long) previously equilibrated with 1× PBS buffer and washed with ten column volumes of 1× PBS (1.0 mL/min).

The flowthrough was discarded and AtCCD1-GST was eluted with 10 mL of the elution buffer

(50 mM Tris, 15 mM reduced glutathione, pH 8.0) at 4°C. Protein elution was monitored by UV absorbance at 280 nm. The protein was concentrated using 10 kDa Amicon tubes and then the buffer was exchanged with 1× PBS containing 0.1% Triton X-100.56 This allows for the protein to retain activity for up to one month at 4°C. The concentration was determined using a bicinchoninic acid assay and the purity of the protein was confirmed using SDS-PAGE (10% gel).

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Regeneration of GST column

The glutathione resin was generated by washing with 20 mL of buffer A (0.1 M Tris-Cl,

0.5 M NaCl, pH 8.5) and the same volume of buffer B (0.1 M sodium acetate, 0.5 M NaCl, pH

4.5). The column was then stored in 20% ethanol at 4°C.

Amylose affinity purification

The crude lysate was collected after centrifugation and added to the amylose column (1 cm i.d.; 18 cm long) equilibrated with 20 mM Tris, 2 mM NaCl, pH 7.4. The manufacturer’s procedure was followed. The column was washed with ten column volumes (0.3 mL/min) 20 mM Tris-HCl, 0.2 M NaCl, pH 7.4. The flowthrough was discarded and AtCCD1-MBP was eluted using 10 mM maltose in 10 mL 20 mM Tris-HCl, 0.2 M NaCl, pH 7.4. Protein elution was monitored by UV absorbance at 280 nm. The protein was concentrated using 10 kDa

Amicon tubes and stored in aliquots at -20°C. The concentration was determined using a bicinchoninic acid assay and the purity of the protein was confirmed using SDS-PAGE (10% gel).

Regeneration of amylose column

The resin was washed with 3 column volumes of deionized water followed by the same volume of 0.1% SDS. The column was then washed with 1 column volume of deionized water followed by 5 volumes of equilibrium buffer (20 mM Tris-HCl, 0.2 M NaCl pH 7.4). The column was stored in 20% ethanol at 4°C.

Sephacryl S-200 gel filtration chromatography

The column (1 cm i.d.; 18 cm long) was packed via gravity and washed with one column volume of 1 x PBS buffer (pH 7.4). The crude lysate was added to the top of the column followed by buffer once it reached the top of the resin. Seventy 2 mL fractions were collected (a total of one column volume). The column was then washed with one column volume of buffer. A

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bicinchoninic acid assay was used to determine which fractions included protein and an SDS-

PAGE (10% gel) was used to determine which contained the AtCCD1-MBP. The column was stored in 20% ethanol or in buffer containing 0.04% sodium azide.

Bradford and bicinchoninic assays

These protein concentration determination assay protocols were followed according to the manufacturer’s instructions.

Enzyme Reactions

ZmCCD1 reactions

A stock solution of linoleic acid (Sigma) was prepared by dissolving 1 μL in 900 μL of deionized water. Assays were performed in a total volume of 1 mL containing 50 mM NaPO4

(pH 7.5), 300 mM NaCl, 5 μM FeSO4, 5 mM ascorbate, 20% methanol, and 5 mM linoleic acid.

Deionized water was added to bring the total volume to 900 μL and the reactions were allowed to equilibrate to room temperature. Finally, 100 μL of crude lysate containing ZmCCD1 was added. The reactions were allowed to proceed overnight at 30°C on a rotisserie. They were extracted with an equal amount of EtOAc and analyzed on GC/MS using JON_METH.

AtCCD1 reactions

A microcentrifuge tube contained 5 mM linoleic acid solution, 50 mM NaPO4 (pH 7.2),

300 mM NaCl, 5 µM FeSO4, 5 mM ascorbate, and 100 µL of purified AtCCD1 with sufficient milli Q water to bring the total volume to 1 mL. The reactions were vortexed briefly and put on the rotisserie (LabQuake) to react overnight in an incubator set at 27°C. Immediately before analysis, 1 mM of d12-hexanal was added. The reactions were extracted twice with 0.5 mL portions of EtOAc and the combined organic layers were analyzed by GC/MS using

JON_METH.38

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Autoxidation Trials

To prevent autoxidation, neat α-cyclodextrin (0.39 g) was evaluated. A 4:1 ratio

(cyclodextrin:linoleic acid) was combined, dissolved in 150 μL of water and put into a 70°C water bath for two hours to form the inclusion complex. The paste was dried overnight at room temperature and placed under vacuum for 2 hours. For each reaction, 50 mg of the paste was used. In addition, trials were done to try and enhance the auto-oxidation of linoleic acid to eliminate the use of enzymes. This was done by varying the concentration of FeSO4 and ascorbate, and also including aluminum foil in the reaction system.

Lastly, riboflavin was used as a substitute for the CCD1 enzymes. A 5 mM linoleic acid solution was prepared according to previously described methods in a crimp top GC vial. A small stir bar was added along with 1 mg riboflavin and the vial was capped. Light from a projector was used as the light source and pointed directly at the reaction. It was allowed to react for 24 hours and then subjected to SPME analysis with d12-n-hexanal as an internal standard.

Biphasic reactions

To the previously mentioned reactions, several organic solvents were used as a second phase to extract the hexanal. To each reaction, 0.5 mL of the individual solvent was added. The solvents evaluated were hexanes, methyl tert-butyl ether (MTBE), limonene, chloroform, and ethyl acetate.

GC/MS analysis: JON_METH

Gas chromatography-mass spectrometry was performed on a Hewlett-Packard (HP) 5890

Series II Plus gas chromatograph (He carrier gas; 1.0 mL/min; split/splitless injector 220ºC; injection volume 1µL) with a DB-17 column (30m long; 250 µm i.d; 0.25 μm thickness). The initial temperature was set to 60°C for 2 minutes (2 minute solvent delay). The temperature was

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programmed from 60ºC at 10ºC/minute to 250ºC (hold for 10 minutes). The instrument was equipped with an auto-sampler. The GC was coupled to a HP 5971 series mass selective detector. n-Hexanal (Aldrich 66-25-1), d12-n-hexanal (CDN Isotopes D-6265), and linoleic acid

(ACROS 60-33-3) retention times were determined using standard solutions and verified by comparison with standards from the NIST library.

Site Saturation Mutagenesis96

For each mutagenesis position, a set of primers were designed for each of the amino acids to be evaluated. Each changed a single codon and took the E. coli codon bias into account. The forward and reverse primers were allowed to overlap at the position desired for mutagenesis and had Tm values between 68-70°C. PCR amplification was performed by combining 5× Phusion

HF Buffer (10 µL), purified pEA1/pEA2 template (~1 ng), forward and reverse primers (0.5 µM each), dNTP mix (200 µM each), and finally 5× Phusion Hot Start II High Fidelity DNA

Polymerase (2 U). The PCR program parameters were as follows: 98°C for 30 seconds followed by 25 rounds of 98°C for 10 seconds, a gradient of 62-72°C for 30 seconds, 72°C for 3 minutes and 30 seconds. Lastly, a final extension was done at 72°C for 7 minutes and 30 seconds followed by a hold at 4°C until removal from the instrument.

Each PCR product was purified using the Wizard PCR Clean-Up Kit using the manufacturer’s procedure and the DNAs were eluted with a final volume of 50 µL sterile water.

The DNAs were then digested with DpnI, purified once again with the same PCR clean-up procedure, and eluted with 50 µL of sterile water. Some of purified DNA (1.5 µL) was used to transform the mutants into E. coli Electro Ten Blue cells via electroporation.

For each transformation, a few colonies were selected, the DNA was purified (Wizard

Mini Prep DNA Purification Kit), and the mutation was confirmed by Sanger sequencing. Once

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confirmed, the same transformation procedure was followed, but into electrocompetent E. coli

BL21(DE3)-Gold cells. The new plasmids were then expressed in E. coli and grown in the same fashion the wildtype enzyme (AtCCD1 purification procedure).

Results and Discussion

Linoleic Acid Behavior under Aqueous Conditions- Light Scattering Studies

In our proposed route to n-hexanal using CCDs, linoleic acid must be dispersed into water. Because of its very low solubility under aqueous conditions, we examined its physical state when present at concentrations needed for enzyme-catalyzed cleavage. Various dilutions of linoleic acid were prepared ranging from 0.001 to 0.05%, making sure that the samples remained non viscous. Each was analyzed by light-scattering in collaboration with the Savin group

(University of Florida).

Once the light scattering experiments were completed, the angle dependence was calculated (q2) according to the following equation by the instrument:

q=(4 π no)/λ * sin (θ/2)

Here, no is the refractive index, λ is the incident laser wavelength, and θ is the angle at which the detector is located with respect to the sample cell. The decay rate was then found through the following equation:

2 Γ= q * Dt

2 Here, Dt is the translational diffusion coefficient with respect to the angle dependence (q ) and Γ is the decay rate. Finally, the decay rate was plotted against the angle dependence to determine if there was an actual angular dependence. Small spherical particles will result in a horizontal line, thus having no angular dependence. Particles with a shape other than a sphere will show an angular dependence, which would be consistent with aggregation.97 The slopes of these plots

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yield Dt, which is further used in the Stokes-Einstein equation to calculate the radius of the hydrophobic spheres.

The results of this study are shown in Figure 2-3a. It was ultimately determined that there is a positive correlation between the concentration of linoleic acid and micelle size (Figure 2-3a), which was expected due to the lack of aqueous solubility. The micelle size of linoleic acid in our samples was between 60 and 90 nm.

A B

C

Figure 2-3. Light scattering experiment results A) of linoleic acid dilutions, B) AtCCD1 without any detergent added, and C) AtCCD1 with 0.1% triton X-100.

We were also interested in the possibility that AtCCD1 might also show aggregation under the reaction conditions. To determine this, AtCCD1 concentrations of 0.01 to 0.05% were evaluated. Based on the measurements from the crystallographic structure (PyMol), the radius of

AtCCD1 should be 67.2 Å (6.7 nm). Without the addition of detergent, high levels of enzyme aggregation were occurring (Figure 2-3b). It was determined that the micelle size was between

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21 and 26 nm, which is much higher than expected. The same samples were analyzed in the presence of 0.1% triton X-100, which significantly decreased the observed size to 5 to 6 nm, near the expected for non-aggregated AtCCD1 (Figure 2-3c). This confirmed that detergent was likely necessary for AtCCD1-catalyzed reaction to prevent protein aggregation.

Figure 2-4. Light scattering results mimicking actual enzymatic reaction conditions

The final set of experiments was carried out with both linoleic acid and AtCCD1 present in the sample cells, which mimicked the actual reaction systems. As seen in Figure 2-4, measurements of samples with 0.005% to 0.025% linoleic acid were consistent with a micelle size of 20 nm. The 0.05% linoleic acid showed a decreased micelle potential. Using 5 mM of linoleic acid in the AtCCD1 reactions would therefore lead to lower amounts of micelle formation. This was not expected, but it is thought that linoleic acid may interact with AtCCD1, decreasing its aggregation. In Figure 2-4, the ‘open’ symbols along the top of the graph represent linoleic acid samples without added AtCCD1. Ultimately these results show that in the presence of enzyme, linoleic acid micelles are broken up and are able to interact with the enzyme more

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readily. Lastly, Figure 2-4 re-emphasizes that adding triton X-100 (or any detergent) to the reaction system prevents the aggregation of AtCCD1. n-Hexanal Quantitation

SPME quantitation

Three fibers were evaluated for SPME hexanal recovery: PDMS/DVB, CAR/PDMS, and

PDMS/CAR/DVB. From these experiments, it was determined that the PDMS/CAR/DVB fiber recovered the most n-hexanal from aqueous solutions (Figure 2-5). It is known that SPME concentrates volatile samples for analysis and this was observed in our experiments. Several compounds that were not observed through simple solvent extractions were detected, including

2-butyl-2-octenal, which was later determined to be a byproduct of n-hexanal. PDMS/CAR/DVB fibers are the preferred fiber for odors and flavors, which would describe n-hexanal.88a Our results therefore made sense since this fiber type would be the most efficient for absorbing n- hexanal.

Figure 2-5. n-Hexanal recovery from various SPME fibers based on their GC peak areas

Several temperatures between 40 and 80˚C were evaluated for the equilibration temperature. The equilibration phase allows for volatiles to collect in the headspace of the vials before fiber insertion. In addition, the equilibration time period was evaluated from 0 to 30 minutes. It was determined that 75°C for 30 minutes was the most efficient for the collection of

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volatiles. This time period was similar to that reported by Garcia-Llatas et al.98 The high temperature could disturb the fiber matrix and shorten its lifetime, but we did not experience any change in n-hexanal absorption throughout our experiments. Methyl benzoate was used as an internal standard to quantitate the adsorbed n-hexanal. The extraction time was evaluated from 0 to 60 minutes. The extraction phase occurs once the SPME fiber is inserted into the headspace of the vial for volatile absorption. It was determined that a 30 minute time period was the most efficient for n-hexanal absorption; longer extraction times yielded decreased absorption. This extraction time frame was also determined to be the most efficient for n-hexanal by Garcia-

Llatas et al.98

A standard curve was prepared for the absorption of n-hexanal with the

PDMS/CAR/DVB fiber using the optimum conditions previously described. It was determined that the fiber was becoming saturated with n-hexanal; it was therefore not possible to prepare a usable standard curve with this method. This showed that SPME is very sensitive, which was preferable for detecting the low concentrations of n-hexanal. Unfortunately, when the optimized

SPME method was used in conjugation with our CCD1-catalyzed reactions, it was found that the data were not reproducible. We therefore sought a different method to quantitate n-hexanal produced by biocatalytic routes.

Other methods for extracting and detecting n-hexanal

The DNPH derivitization procedure was applied to a sample of pure n-hexanal. Based on the melting point and NMR data, the desired derivative was produced. Unfortunately, when this technique applied to our enzymatic reactions, no DNP-derivative of n-hexanal was observed.

This could have been due to low concentrations of n-hexanal produced or something in the reaction system was preventing the derivitization.

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Since n-hexanal has low solubility in water, several biphasic systems were evaluated to extract it as soon as it was made enzymatically. Several organic solvents were tested including:

CHCl3, hexanes, octane, CH2Cl2, and EtOAc. None of the biphasic systems resulted in n-hexanal formation in the presence of CCD1 enzymes. Since linoleic acid also has low water solubility, we suspected that it was also extracted into the organic phase, so that it could not interact with the CCD1 enzyme in the aqueous phase. In addition, it was observed that when enzyme was included, it would precipitate from the solution when several of the organic solvents were present.

Several solid phase extraction techniques were also tested in order to remove n-hexanal before it could react further. These included XAD4, XAD7, and (aminomethyl) polystyrene.

XAD4 is used most commonly to remove small, hydrophobic organic molecules. XAD7 is used most commonly to remove molecules with molecular weights up to 60,000.99 Both were evaluated for their ability to recover n-hexanal; XAD7 was more efficient. XAD7 was also able to recover n-hexanal from enzymatic reactions whereas XAD4 was not. The optimal pH for this removal system was a pH of 6. The main drawback was that the maximum recovery of n-hexanal using XAD7 was 30%, so a more efficient removal method was needed.

We also tested (aminomethyl) polystyrene for recovering n-hexanal. This resin includes a primary amine, which could form Schiff’s base linkages with n-hexanal. This reaction is reversible, which should allow n-hexanal to be easily recovered. When added to the enzymatic reactions, the resin actually catalyzed an aldol condensation of n-hexanal. The main product recovered was 2-butyl-2-octenal, which could not be converted back into n-hexanal. The biphasic and resin systems were ultimately deemed unsuitable for n-hexanal recovery.

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We therefore chose simple solvent extraction as a way to quantitate and recover n- hexanal from enzymatic reactions. Since n-hexanal was detectable by GC/MS, we decided to use d12-n-hexanal as an internal standard since it should have the same properties as n-hexanal. Both normal and deuterated n-hexanal were detected by GC/MS, with baseline separation and in the expected 1:1 ratio (Figure 2-6). The major drawback is that n-hexanal easily decomposes, and d12-n-hexanal did as well. Its high cost and short shelf life prompted us to use lauric acid or methyl benzoate as internal standards since neither of these compounds interferes with the other peaks in the chromatogram.

Figure 2-6. GC of d12-n-hexanal and n-hexanal separation

Linoleic Acid Quantitation

It was determined that linoleic acid could not be observed by GC without prior derivatization. The first method used for this purpose was methyl ester formation. This procedure required 24 hours and used high concentrations of HCl.91 We therefore explored a silylation procedure using N-methyl-N-(trimethylsilyl) trifluoroacetamide (MSTFA). This procedure took

30 minutes, although it required the use of distilled pyridine.76, 100 Both of these procedures were successful in allowing linoleic acid quantitation by GC/MS.

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Cloning and Expression of AtCCD1

The original ZmCCD1 overexpression strain gave relatively low protein yield, which also resulted in very poor yields (~13%) of n-hexanal from linoleic acid. The levels observed were nearly the same as those from the autoxidation of linoleic acid.

We therefore turned to AtCCD1 as an alternative that could be produced in E. coli at higher levels and was better characterized.36 The AtCCD1 gene was chemically synthesized by

GenScript with flanking NdeI and XhoI sites at the 5’- and 3’- ends, respectively, and inserted into pUC57 (Figure 2-7). The AtCCD1 gene was fused at its N-terminus to GST by subcloning the NdeI, XhoI fragment between these sites on pBS2. The fusion protein is under the control of a T7 promoter in the resulting plasmid, designated pEA1.

Figure 2-7. Construction of pEA1 from pBS2 and pUC57-AtCCD1

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After verifying the structure of pEA1 by sequencing, it was used to transform E. coli strain BL21(DE3)-Gold. After induction, the crude lysate was applied to a glutathione resin and the desired fusion protein was eluted by adding reduced glutathione. Two prominent bands were observed by SDS-PAGE (Figure 2-8). The more prominent band around 87 kDa is consistent with the desired GST-AtCCD1 fusion protein while the band around 26 kDa is likely to be free gluathione S-transferase.

Figure 2-8. 10% SDS-PAGE of the purified GST-AtCCD1 fusion protein using affinity chromatography

While this expression system produced soluble fusion protein that could be purified, it only yielded 24-50 μg of fusion protein from a 0.5 L culture. It also showed low catalytic activity when assayed against β-apo-8’-carotenal. These results were contrary to those found by Vogel et al. who reported that the GST tag aided in protein solubility and activity.38 This was not observed in our study.

When 5 mM linoleic acid was tested as a substrate for the GST-AtCCD1 fusion protein, control reactions lacking protein actually produced greater amounts of n-hexanal than those with the fusion protein: no protein, 0.49 mM (10% yield); GST-AtCCD1, 0.32 mM (6% yield). In a

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few trials, those with the fusion protein produced more n-hexanal; however, the levels were within experimental error of those from the negative controls.

We suspected that reduced glutathione, the GST-affinity tag or the GST-AtCCD1 fusion protein was interfering with the enzyme-catalyzed cleavage of linoleic acid. Reduced glutathione was tested in a recovery experiment with n-hexanal and it was determined not to interfere. The contaminating glutathione S-transferase was removed by ultrafiltration using Amicon tubes with a molecular weight cutoff of 60kDa and was determined not to interfere. The unrelated GST- fusion protein encoded by pBS2 protein (Figure 2-7) was overexpressed and purified in the same manner as the GST-AtCCD1 as a control to determine whether the GST-tag interfered with n- hexanal production and/or recovery. The GST-OYE 2.6 fusion protein was added to an n- hexanal recovery experiment. Solvent extraction revealed no n-hexanal, consistent with the notion the GST-tag was interacting with the n-hexanal, resulting in no product recovery. This led us to investigate other affinity tags for AtCCD1. This is in correlation with what was observed by

Vogel et al. that the GST fusion could interfere with enzyme activity.

Figure 2-9. The construction of pEA2 from pMAL-c5x and pEA1

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The second affinity tag selected was maltose binding protein (MBP). This tag was expected to increase the solubility of the fusion protein and it also has a cleavage site for a factor

Xa protease at the fusion junction. This plasmid was synthesized according to Figure 2-9.

When MBP-AtCCD1 was overexpressed, a large amount of the protein was synthesized as inclusion bodies or misfolded proteins. A temperature and time study was done for MBP-

AtCCD1 expression at 18, 25, 30, and 37˚C and for a 24 hour time period. It was determined that

18˚C with overnight expression was optimum and solved the inclusion body problem. A third plasmid was also made to express AtCCD1 without a tag using pET22b. A SDS-PAGE (10% gel) was run as a comparison of the expression levels of each CCD1 expression system developed (Figure 2-10). As seen in Figure 2-10, MBP-AtCCD1 had the best expression level and resulted in about 350 mg of protein per liter of culture. The optimal IPTG concentration, induction temperature, and induction time after IPTG feeding were 0.4 mM IPTG, 18°C and 18 hours.

Figure 2-10. Expression level comparison a) pEA1, b) pEA2, c) pEA3, and d) ZmCCD1

MBP-AtCCD1 Cleavage of Linoleic Acid

CCD1 has not been previously known to cleave any substrate other than carotenoid compounds. Utilizing the reaction conditions reported by Vogel et al., there was little enzymatic

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activity with linoleic acid.38 Linoleic acid is known to be susceptible to autoxidation. Reactions without enzyme were therefore necessary to determine how much cleavage was due to enzymatic activity. When compared to each other, the n-hexanal production was similar or within an error factor (Figure 2-11).

Figure 2-11. n-Hexanal production from enzymatic and non-enzymatic reactions

Effects of pH, temperature and buffer on linoleic acid cleavage into n-hexanal by MBP- AtCCD1

Since FeSO4 and ascorbic acid are radical initiators, they lead to enhanced autoxidation of linoleic acid. When these components were omitted from the reactions, no n-hexanal production was observed. They were, however, necessary for any enzymatic activity. To enhance the cleavage of linoleic acid with MBP-AtCCD1, several parameters were varied including buffer, pH and temperature. When various buffer systems were evaluated, it was determined that those with higher pKa values resulted in more linoleic acid cleavage (Figure 2-12a). The reaction pH was evaluated from 3.5 to 8 and it was determined that MBP-AtCCD1 has optimum activity at pH 6 (Figure 2-12b). Without enzyme, the recovery of n-hexanal remained constant at each pH value investigated. Finally, the temperature was varied from 4 to 37°C and it was determined that 25°C was optimum (Figure 2-12c). All further experiments were therefore performed at pH

6, 25°C, and in 50 mM sodium phosphate buffer.

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Figure 2-12. Comparison of various changes to MBP-AtCCD1 reaction parameters A) the effect of various buffers on the cleavage of linoleic acid by MBP-AtCCD1, B) pH effects on n-hexanal recovery with and without MBP-AtCCD1 and C) effect of temperature on linoleic acid cleavage

Effects of CLEA and acetylation on the MBP-AtCCD1 cleavage activity of linoleic acid

In experiments using crude lysates containing MBP-AtCCD1, it was found that the recovery of n-hexanal decreased drastically. We suspected that surface Lys residues might form

Schiff’s bases with n-hexanal and thereby diminish its recovery. MBP-AtCCD1 was therefore acetylated in an attempt to prevent these interactions. The acetylation procedure worked efficiently and easily. Reaction 4 resulted in 60 acetylation sites and used acetic anhydride and sodium acetate. The use of sodium acetate alone resulted in no acetylation and the use of acetic anhydride alone resulted in 17 acetylated sites of MBP-AtCCD1. There are 78 possible acetylation sites in MBP-AtCCD1 and reaction 4 ensures that the most reactive were derivitized.

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Enzymes acetylated using reaction 3 had the most activity with β-apo-8’-carotenal. We believe this resulted from acetylating only the most active residues in the enzymes, which resulted in less structural changes. Since no improvement in n-hexanal production was observed, further efforts in this area were halted.

We also briefly explored the use of a cross-linked enzyme aggregate (CLEA) that included MBP-AtCCD1 since this also involved reacting the surface lysines with glutaraldehyde.

While the optimum (NH4)2SO4 saturation levels were determined (80% for crude lysate and 60% for purified enzyme), after adding the glutaraldehyde to these solutions all activity was lost in the

β-apo-8’-carotenal reaction. Unfortunately, all CLEAs produced also had no catalytic activity with MBP-AtCCD1.

Purification of MBP-AtCCD1 for enzymatic reactions

To avoid losing n-hexanal to side-reactions from crude lysates, we decided to use purified

MBP-AtCCD1 since this resulted in almost 100% n-hexanal recovery in control extractions. The fusion protein was purified using an amylose column, since the maltose binding protein shows affinity towards amylose. The amylose column could only be regenerated up to 5 times and it was found that it could not be used with an FPLC due to collapse at higher pressures. In addition, most of the fusion protein failed to bind to the resin and a large amount was observed in the column flow through (Figure 2-13).

Figure 2-13. Purification of AtCCD1-MBP with amylose column a) 10x diluted purified, b) 2x diluted purified and c) column flow through

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A Sephacryl gel filtration column (size exclusion) was therefore used for purification at room temperature. This procedure was much simpler and resulted in a more pure protein with decreased MBP isolation (Figure 2-14). Ultimately, the Sephacryl column was preferred for

MBP-AtCCD1 purification, because it could also be used at room temperature with a peristaltic pump.

Figure 2-14. Comparison of MBP-AtCCD1 purification a) Sephacryl S-200 (10x dilution) and b) amylose (2x dilution)

Effects of alcohol in the reaction mixtures to solubilize linoleic acid

To increase the solubility of linoleic acid, it was first dissolved in EtOH before adding it to the reaction system. It was determined that making the linoleic acid stock in this fashion resulted in more cleavage of linoleic acid. Concentrations of ethanol up to 5% in the enzymatic reactions were tolerated; above that, the recovery of n-hexanal decreased (Figure 2-15). In non- enzymatic control reactions, it appeared that EtOH concentrations above 2% resulted in lower n- hexanal recoveries. In the original reaction conditions developed by Vogel et al., 20% MeOH was included.38 It was discovered that for linoleic acid, including MeOH in the reactions caused a decrease in n-hexanal production. We therefore deleted MeOH from reaction mixtures and used EtOH to make the linoleic acid stock solution.

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Figure 2-15. Effects of increasing the concentration of EtOH in the reaction systems

Autoxidation of Linoleic Acid

Linoleic acid is prone to autoxidation and n-hexanal is one of the known products from this process.101 Because radical-producing conditions would also be found in CCD1-catalyzed reactions, it was important to determine the background levels of n-hexanal production by non- enzymatic pathways prior to investigating the contributions of CCDs.

It was previously observed in our studies that higher levels of FeSO4 and ascorbate lead

101 to increased levels of linoleic acid autoxidation. The FeSO4 and ascorbate concentrations were therefore varied from 0-50 μM and 0-50 mM, respectively (Figure 2-16). The n-hexanal concentration increased slightly when increased simultaneously; however, the overall amount was below 0.25 mM (0.025 mg, 5% yield). Linoleic acid autoxidation was also compared at

25˚C versus 37˚C over 3 days. Less autoxidation was observed at the higher temperature (0.15 mg, 3% yield). At 25˚C, n-hexanal production increased until 48 hours, at which point it plateaued. The difference in autoxidation is believed to be due to linoleic acid being more soluble at higher temperatures.

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A B

C

Figure 2-16. Autoxidation of linoleic acid compared to enzymatic reactions a) increasing FeSO4, b) increasing ascorbate, and c) increasing FeSO4 and ascorbate simultaneously

Riboflavin was evaluated as a simpler replacement for the CCDs since this substance is known to generate a singlet oxygen species that cleaves alkenes into aldehydes under the influence of high visible light fluxes.102 Trace amounts of n-hexanal were observed in illuminated reactions with no riboflavin and also when riboflavin was present, but lacked illumination. Yields of up to 0.15 mg (3% yield) n-hexanal were observed in riboflavin-catalyzed reactions under constant visible light from a commercial slide projector. Unfortunately, because this concentration of n-hexanal was lower than those produced through enzymatic reactions or autoxidation trials, further trials with riboflavin were discontinued.

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In an attempt to prevent autoxidation of linoleic acid during enzyme-catalyzed reactions,

α-cyclodextrin was included to capture the linoleic acid. While this was successful in eliminating autoxidation, it also appeared to block the cleavage of linoleic acid by MBP-AtCCD1since no n- hexanal was observed.

We also tried to use catalase to prevent spontaneous oxidation of linoleic acid, which will decompose hydrogen peroxide. Interestingly, when catalase was included in the reactions, it led to greater n-hexanal production resulting in 34% yields (Figure 2-17).

Figure 2-17. Effects of adding catalase to the reaction systems with and without MBP-AtCCD1

Other AtCCD1 Substrates

Unfortunately, linoleic acid itself yielded only very low concentrations of n-hexanal production using MBP-AtCCD1. Several other potential substrates were also tested, including conjugated linoleic acid isomers, palmitoleic acid, palmitic acid, and β-apo-8’-carotenal.

Palmitoleic acid contains a single alkene at the C9 position while palmitic acid is a negative control with no unsaturation. Neither of these substrates yielded short-chain aldehydes in the presence of MBP-AtCCD1. This was expected since neither structure resembles a substrate known to be accepted by AtCCD1. They were chosen because they should not have been cleaved by the enzyme. There are four conjugated linoleic acid (CLA) isomers: 9Z,11E CLA, linoeladic

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acid, 9E,11E CLA, and 10E,12Z CLA. The 9Z,11E CLA isomer would be expected to lead to n- heptanal instead of n-hexanal due to the change in alkene position. It is worth mentioning that

9E,11E CLA was insoluble in the reaction mixtures. The CLA isomer conversion data can be observed in Table 2-2.

Table 2-2. Conversion of CLA isomers into hexanal

Compound Structure No Enzyme AtCCD1

Linoeladic Acid 0.080 mM 0 mM

9E,11E CLA 0 mM 0 mM

10E,12Z CLA 0.170 mM 0.949 mM

β-Apo-8’-carotenal is a synthetic substrate that is known to be cleaved by the AtCCD1 enzyme. Cleavage was not achieved with the crude lysate, however, when the purified protein was used, almost 100% conversion occurred in 1 hour. This is contrary to the results of Vogel et al. and Schilling et al., who reported that the purified protein gave lower activity.38, 56 This substrate was used to test the enzyme activity each time it was purified since the reactions were quick and led to consistently high conversion rates.

Ligand Binding Studies of MBP-AtCCD1

As mentioned previously, β-apo-8’-carotenal is a synthetic substrate cleaved by AtCCD1 and it is the only other known substrate besides carotenoid compounds. Here, it was used to confirm the activity of AtCCD1 fusion proteins. Upon substrate cleavage, the color changes from orange to yellow within 1 hour at 25°C. When analyzed via HPLC, 90% conversion was reached within a 1 hour time period (Figure 2-18).

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Figure 2-18. HPLC data for β-apo-8’-carotenal reactions a) no AtCCD1 added and b) with AtCCD1

To determine whether the lack of n-hexanal production by AtCCD1 was due to issues with substrate binding, this system was used in inhibition studies with various fatty acids at concentrations of 1 mM. The materials tested for inhibition are listed in Table 2-3.

Table 2-3. Inhibition trial results for various materials evaluated with MBP-AtCCD1

Complete inhibition Partial inhibition Complete conversion Linoleic Acid 9Z, 11E CLA β-apo-8’-carotenal Methyl Lineolate Linoeladic Acid Eladic Acid 10E, 12Z CLA Arachidonic Acid Trans-Crotonic Acid Linolenic Acid Palmitoleic Acid Hexanoic Acid Oleic Acid Valeric Acid E,E-2,4-Decadienal 1-Octene Erucic Acid Triglyceride of Linoleic Acid Lauric Acid E-4-Decenal n-Hexanal n-Heptanal *Structures available in Appendix B

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When analyzing the results from Table 2-3, it was discovered that linoleic acid actually led to a complete inhibition of enzymatic activity. We believe this was due to linoleic acid having two cis-alkene bonds, resulting in a more kinked system as compared to the carotenoid compounds. Carotenoids are all-trans compounds and the scissile bond is isomerized by AtCCD1 prior to cleavage. The kinked system could prevent linoleic acid from entering the active site tunnel of the enzyme. It was later determined that AtCCD1 accepted 9-cis-violaxanthin as a substrate rendering this theory unlikely. It is also possible that linoleic acid can enter the active site of AtCCD1; however an alkene bond is not correctly positioned for cleavage, near the catalytic Fe within the active site. The second possibility is more likely since there was no evidence of β-apo-8’-carotenal cleavage in reactions that included linoleic acid. The results of this inhibition study ultimately led to the conclusion that linoleic acid is not a good substrate for

AtCCD1 cleavage, likely acting as a non-competitive inhibitor. Kinetic studies were not performed to validate the type of inhibition. It was also determined that n-hexanal slightly inhibited catalytic activity. Together, these results showed that further studies with wild-type

AtCCD1 and linoleic acid were likely to be unsuccessful.

The all-trans isomer of linoleic acid was tested in this same system and was found to be insoluble under the reaction conditions. Since AtCCD1 is known to act as an isomerase prior to compound cleavage, the 9Z, 11E conjugated linoleic acid isomer was examined in the inhibition reaction. The desired cleavage position is in the correct trans-orientation for cleavage; it was therefore thought that AtCCD1 would accept it more readily. The inhibition level decreased with this isomer drastically, so it seemed as if it would be a good substrate for testing. Unfortunately, this isomer was extremely expensive and our industrial partner required that we use linoleic acid

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itself. Known isomerase enzymes can accomplish this transformation; however, the conjugated linoleic acid isomers would further need separated from this process.

The other compounds evaluated under these conditions were tested to evaluate the extent of AtCCD1 substrate scope. They included several variations of different fatty acids and small aldehyde compounds. The structures of the compounds evaluated can be found in Appendix B.

In general, the smaller compounds did not interfere in carotenal cleavage, while the larger compounds had a mixture of slight to complete inhibition. This was expected, since AtCCD1 accepts long chain and non-polar substrates.

Homology Model of AtCCD1

Crystal structures of membrane associated proteins are difficult to determine, so we therefore made a homology model of AtCCD1 based on a crystal structure from a related protein in the same family (VP14).103 This model was predicted using the I-TASSER server developed by the Zhang lab at the University of Michigan.104 It was predicted with a C-score of 0.89, which is a measure of the confidence in the quality of the model. These scores typically range from -5 to 2 and the higher the score, the greater the confidence in the predicted model. The dome region of the enzymes (Figure 2-19) varies considerably, leading to a lower C-score value. A score of

0.89 is a relatively good score and confidence can be put into the predicted structure seen in

Figure 2-16. Similar to other closely related enzymes (ACO1 and VP14), AtCCD1 is likely comprised of a 7-bladed propeller made up of antiparallel β-sheets with a hydrophobic tunnel in which the substrate cleavage occurs. On top of the stable propeller is a dome region and an exit from the substrate tunnel allowing for product expulsion. Figure 2-16 also shows the proposed position of the substrate bound in the active site tunnel in purple (3-hydroxy-8’-carotenal).

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Figure 2-19. Homology model of AtCCD1 with bound substrate predicted by I-TASSER

Similar to other enzymes in this family, Fe is coordinated by 4 histidine residues within the active site tunnel for cleavage. This coordination can be observed in Figure 2-20a. In addition, linoleic acid was modelled into the system and the coordination to the Fe can be seen in

Figure 2-20b. The alkene to be cleaved aligns fairly close to the catalytic Fe; however, its angle is slightly off. Since we are making this assumption based on a homology model and manually adding linoleic acid, it was determined that the angle was close enough to attempt using this enzyme for linoleic acid cleavage. It is worth mentioning that the program used for predicting the enzyme structure was not able to add molecular oxygen into the coordination scheme. The catalytic Fe has six positions open for coordination: four histidine residues, molecular oxygen, and a water molecule (Figure 2-20a).41 The molecular oxygen is held close to the Fe atom by two additional amino acids: threonine 175 and phenylalanine 402.

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Figure 2-20. Catalytic Fe coordination within the active site tunnel of AtCCD1 A) the amino acid residues responsible for coordination and B) linoleic acid coordination to this system

In addition to modeling the Fe coordination, linoleic acid itself was modeled into the active site tunnel. The position of linoleic acid was determined by overlapping it with a known substrate that was included in the predicted structure. This overlap can be observed in Figure 2-

21a. For clarity purposes, the rest of the protein was not shown in order to highlight the overlap of the key alkenes (Figure 2-21b). While the double bonds and half of the linoleic acid structure overlaps well with 3-hydroxy-8’-carotenal, the methyl terminal end is significantly different

(Figure 2-21a). It seems likely that this discrepancy contributes to the inability of AtCCD1 to cleave linoleic acid efficiently.

Figure 2-21. Overlap of 3-hydroxy-8’-carotenal with linoleic acid A) within the active site tunnel and B) to show the overlap of the alkene to be cleaved

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AtCCD1 Mutagenesis studies

In addition to the I10V mutation initially made to AtCCD1 that increased its overexpression yield, it was of interest to mutate specific positions within the active site tunnel to better accommodate linoleic acid. The homology model was able to predict the amino acids that help to position the substrates near the Fe and oxygen. These include Phe 98, Ile 140, Gly 141,

Thr 175, Gln 188, Glu 189, Ala 190, Pro 267, Met 269, His 271, Phe 336, Asp 401, and Phe 402.

Each of these residues is represented by green spheres at the alpha carbon of the amino acid in

Figure 2-22. They surround the modeled linoleic acid.

Figure 2-22. Residues respoonsible for positioning linoleic acid near the catalytic Fe

Each residue was evaluated for possible mutagenesis and Phe 98 and Met 269 were chosen for initial studies.

The first residue evaluated was Met 269. It was chosen due to the observed steric clash with linoleic acid in the homology model (Figure 2-23a). By changing this amino acid to a smaller one (such as Ala), we hoped to relieve this clash and allow linoleic acid to be better accepted by AtCCD1. Alanine is also a less polar residue than methionine, so it would interact to a better degree with the nonpolar tail of the fatty acid. Met 269 is located at the entrance to the

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substrate tunnel and is exposed to the aqueous environment, so it could also be preventing linoleic acid from entering the tunnel.

The second residue chosen was Phe 98. It also showed a steric clash with linoleic acid, although it is located further into the active site (Figure 2-23b). It was thought that by changing this residue to a smaller one (such as Ala), linoleic acid would be better accommodated in the

AtCCD1 active site. In addition, Phe 98 lies near the catalytic Fe site. If linoleic acid is too close to the Fe, alkene cleavage may be inhibited. It was also of interest to combine the two mutations together to observe their combined effects on linoleic acid acceptance by AtCCD1.

Figure 2-23. The two positions proposed for mutagenesis studies A) Met 269 and B) Phe 98

Each of these residues was mutated to an alanine in pEA1 and the expression levels were significantly lower than with the wild type enzyme. The mutations were therefore transferred into pEA2 and the expression levels greatly increased with the M269A mutant, but stayed the same for the F98A mutant. The mutants were analyzed in the β-apo-8’-carotenal assay for protein activity. It was observed that M269A has roughly 50% the activity than the wild type enzyme. The F98A mutant showed no activity and it is believed to be due to low expression levels. The same was observed with the F98A, M269A mutant. We believe that making changes to the inner substrate tunnel leads to decreased protein stability, since the mutants with F98A

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showed little protein expression. Further studies would need to be done to confirm this hypothesis. Given these disappointing results, these efforts were halted.

Conclusion

The maximum production of n-hexanal from CCD1-mediated cleavage of linoleic acid was around 0.5 mM (0.05 mg, 10% yield). Despite several attempts to increase the enzymatic activity, it was ultimately determined that the AtCCD1 was not efficient at cleaving linoleic acid into n-hexanal. To be a viable industrial process, much higher n-hexanal concentrations would be required. Autoxidation trials lead to a maximum n-hexanal concentration of around 0.7 mM

(0.07 mg, 14% yield) consistently. While this was an improvement over enzyme-catalyzed reactions, we were unable to exceed this concentration despite numerous attempts.

Based on inhibition studies, it was determined that AtCCD1 binds linoleic acid, but does not accept it as a substrate. It seems likely that the alkene alignment with the catalytic Fe is not correct for cleavage. The conjugated linoleic acid isomer (9Z, 11E) showed promise as a substrate; however, due to its high cost only a few trials could be done. The inhibition trials with this substrate showed reduced cleavage of β-apo-8’-carotenal, but not complete inhibition.

Making the conjugated linoleic acid isomer would require the use of an isomerase followed by

AtCCD1, which was not a desirable method.

When the homology model was created and evaluated, it showed that linoleic acid’s alignment with the catalytic Fe was slightly off, which supports our hypothesis. Linoleic acid also sticks out more into the active site tunnel than known substrates, which could alter the internal structure of the enzyme. Mutations were made to try and better accommodate linoleic acid in the active site. These led to no or decreased enzyme activity.

Ultimately this CCD1 route did not produce enough n-hexanal from linoleic acid and we therefore turned to alternative strategies that are discussed in future chapters.

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CHAPTER 3 PRODUCTION OF 13S-HYDROPEROXY-9Z, 11E-OCTADECADIENOIC ACID USING LIPOXYGENASE (LOX1) ISOLATED FROM SOYBEANS

Background

In the previous chapter, we explored the use of carotenoid cleavage dioxygenase enzymes to cleave linoleic acid to make n-hexanal. Despite exhaustive efforts, this route resulted in dismal amounts of n-hexanal production. We therefore turned to lipoxygenase enzymes, which install a hydroperoxy group at the C13 of linoleic acid. This reaction has been explored by several other groups in combination with a lyase enzyme to make n-hexanal.30-31 We decided to explore pyrolysis as an alternative to the use of a hydroperoxide lyase for the C-C bond cleavage step in a procedure similar to that that has been applied to castor oil (Figure 3-1). This chapter will report the results of our studies with lipoxygenase to create n-hexanal by cracking 13- hydroxylated linoleic acid, which mimics the structure of ricinoleic acid found in castor oil. The main difference is that the lipoxygenase-derived fatty acid contains a conjugated diene, rather than the homoallylic alcohol of ricinoleic acid.

Figure 3-1. Experimental strategy of using lipoxygenase to create n-hexanal

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Experimental Strategy

13S-Hydroperoxy-9Z,11E-Octadecadienoic Acid Cracking

Trivedi and Vasishtha found that by heating castor oil with rosin, large amounts of n- heptanal were produced.25 In castor oil, the ricinoleic acid is present in its triglyceride form

(triricinolein). The rosin added to the pyrolysis procedure aides in creating free fatty acids. While our proposed procedure would involve the use of free fatty acids (not triglycerides), we still planned to include rosin in the cracking mixture in case it had other beneficial functions during pyrolysis. Before synthesizing C13-hydroperoxy linoleic acid and pyrolyzing this material, we first attempted to reproduce the literature report of castor oil pyrolysis procedure in our lab.

Lipoxygenase

Soybean lipoxygenase is commercially available through various sources. For initial studies, the purified ammonium sulfate suspension was ordered from Sigma; however, when the number and scales of the reactions increased we prepared the enzyme directly from soybeans.

Linoleic Acid Conversion and Reduction

Initial studies using lipoxygenase to convert linoleic acid into 13-hydroperoxy linoleic acid were carried out according to Clapp et al.74 Several reaction parameters were varied to optimize our lipoxygenase reactions.

The lipoxygenase enzyme adds a peroxide group to C13 of linoleic acid. Prior to the thermal cracking procedure, the hydroperoxide must be reduced to the corresponding alcohol.

We used TCEP for this purpose and several trials were run to optimize the TCEP reduction. Had this strategy been more successful in yielding n-hexanal, we would have used glutathione peroxidase as the reductant since this has been shown to be successful.74

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Experimental Methods

Pyrolysis of Castor Oil

Castor oil (Acros, 150 mL) and rosin (MP Biomedicals, 10 g) were mixed in a 250 mL three-neck flask equipped with a distillation head and condenser (Figure 3-2) and heated over a bare flame. After approximately 20 minutes, the distillate began to collect in the receiving flask.

The temperature was kept around 330°C until approximately 50 mL of distillate had collected, then heating was stopped. This is done to avoid forming a yellow, resinous paste in the reaction flask. The water was removed from the distillate by means of a separatory funnel and the residue was dried using anhydrous magnesium sulfate. The distillate was analyzed via GC/MS using

JON_METH.

Figure 3-2. Castor Oil Pyrolysis set up105

GC/MS analysis: JON_METH

Gas chromatography-mass spectrometry was performed on a Hewlett-Packard (HP) 5890

Series II Plus gas chromatograph (He carrier gas; 1.0 mL/min; split/splitless injector 220ºC; injection volume 1µL) with a DB-17 column (30m long; 250 µm i.d; 0.25 μm thickness). The initial temperature was set to 60°C for 2 minutes (2 minute solvent delay). The temperature was programmed to rise from 60ºC at 10ºC/minute to 250ºC (hold for 10 minutes). The instrument

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was equipped with an auto-sampler. The GC was coupled to a HP 5971 series mass selective detector.

Lipoxygenase Reactions

A 0.5 M linoleic acid stock solution was prepared by dissolving 156 μL of linoleic acid in

844 μL EtOH. A 50 mL solution of 50 μM linoleic acid in 50 mM borate, pH 9.0 was prepared from this stock.74 A stir bar was added and the mixture was stirred vigorously for 30 minutes prior to the addition of 0.43 nmol of lipoxygenase (Sigma type 5). For reactions using lyophilized lipoxygenase, 8 mg was added in two additions (second addition at 6 hours). Once the reaction was complete, 29 mg TCEP was added and stirred for 30 minutes. The reaction was acidified to pH 3-3.5 using 1.0 M citric acid. It was then extracted with 30 mL of Et2O followed by two 15 mL extractions with the same solvent. The organic extracts were combined and concentrated on the rotary evaporator. The residue was resuspended in 1 mL hexanes. This procedure was repeated for various concentrations of linoleic acid.

13-(S)-Hydroperoxy-9Z,11E-Octadecadienoic Acid Quantitation

1. Ferrous-xylenol orange (FOX) method

The FOX reagent included 100 μM ferrous sulfate, 100 μM xylenol orange, and 25 mM sulfuric acid dissolved in 9:1 methanol: water. To determine the concentrations of peroxides, 100

μL aliquots of the reaction mixtures were added to 900 μL of the FOX reagent. These were allowed to react at room temperature for 1 hour and then the absorbance was read at 570 nm. A standard curve was prepared with 13-(S)-hydroperoxy-9Z,11E-octadecadienoic acid and the reaction absorbances compared to the known values.74, 106

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2. UV-Vis

Periodically 1 mL aliquots were taken from the lipoxygenase assay and transferred to a cuvette. The UV spectrum was scanned from 200-300 nm and the A234 was monitored for reaction progress. Once the A234 value remained steady, the reaction was considered to be complete. With the higher concentrations of linoleic acid, the aliquots were diluted prior to obtaining a UV spectrum.

3. FAME/MSTFA derivitization

The procedures described in Chapter 2 were used.

Lipoxygenase Isolation from Soybeans

A 50 g sample of whole soybeans was ground into a fine powder using a Waring blender.

They were then defatted through extractions with hexanes until the elute clarified. The defatted soybean meal was extracted with 7.5 volumes of 0.2 M sodium acetate buffer pH 4.5 by slowly stirring for 1 hour. The suspension was filtered through cheesecloth and centrifuged at 6,000 rpm for 20 minutes at 4°C. The supernatant was adjusted to pH 6.8 with 10 M NaOH. All subsequent steps were performed at 4°C. The clarified extract was taken to 30% saturation by slow addition of solid ammonium sulfate and stirred for 1 hour. The suspension was centrifuged at 6,000 rpm for 20 minutes at 4°C. The pellet was discarded and then the ammonium sulfate concentration was slowly raised to 60% saturation. Throughout this process 0.2 M NaOH was added to keep the pH at 6.8. The suspension was centrifuged and the supernatant was discarded. The pellet was resuspended in an equal volume of 20 mM sodium phosphate buffer pH 6.8. Dialysis was carried out against 3 x 2 L 20 mM sodium phosphate buffer pH 6.8, overnight.64 The suspension was centrifuged at 15,000 rpm for 20 minutes at 4°C. Half of the supernatant was stored at 4°C to use directly in reactions. The other half was lyophilized overnight and stored at -20°C.

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13S-Hydroperoxy-9Z,11E-Octadecadienoic Acid Cracking

To a microdistillation flask (Figure 3-3), the lipoxygenase product residue was added along with 2 mL of peanut oil. Finally, a small amount of rosin was added to the flask. The condenser (4°C) was placed into the flask and the residue was submerged into a crystallization dish filled with peanut oil. The dish was filled with peanut oil to closely resemble the temperature inside the microdistillation flask. A thermometer was added to the dish and a

Bunsen burner was lit underneath the oil bath. The temperature was slowly raised to 250°C, which is when the residue bubbling inside the flask ceased and considerable smoking occurred.

The oil was allowed to cool and the condenser was washed with ethyl acetate. The ethyl acetate portions were combined and 1 mM lauric acid was added. The sample was added to a crimp top vial for GC/MS analysis via JON_METH.

Figure 3-3. Microdistillation flask used for cracking procedure and associated condenser

Results and Discussion

Castor Oil Pyrolysis

The procedure developed by Trivedi and Vasishtha was successfully reproduced in our lab.107 The distillate contained three major products: n-heptanal, n-heptanol, and 10-undecenoic acid. These were the expected products and that had been observed in previous studies.108

Although Trivedi and Vasishtha mentioned being able to separate these components with fractional distillation methods, we did not attempt this since our purpose was to determine

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whether we could replicate their results in our laboratory.107 Based on this success, we decided to form the C13 hydroxylated derivative of linoleic acid, which would resemble the ricinoleic acid structure present in castor oil (Figure 3-4).

A) B) Figure 3-4. Fatty Acid Structures A) Ricinoleic acid and B) 13-Hydroxyoctadeca-9Z,11E-dienoic acid

Biotransformation of Linoleic Acid Using Lipoxygenase

Soybean 13-lipoxygenase was chosen to convert linoleic acid into its C13 hydroperoxy derivative. The enzyme was easily obtained, could accept up to 100 mM of substrate and the reaction has been well-studied. We reduced the 13-hydroperoxy moiety to the corresponding alcohol using TCEP. The first trials used Sigma Type 5 soybean lipoxygenase and 50 μM linoleic acid. This reaction was easily monitored by UV-Vis (Figure 3-5a). When the concentration of linoleic acid was increased to 1 mM, the absorbance levels reached their maximum after 20 minutes (Figure 3-5b). While UV-Vis was a useful method for following reactions with relatively low substrate concentrations, a more robust method was needed for higher concentrations.

Figure 3-5. UV-Vis spectrum for lipoxygenase reactions A) with 50 μM linoleic acid and b) with 1 mM linoleic acid

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The FOX method was tested for this reaction system; however, since we used TCEP to reduce the hydroperoxy group before acidifying the reaction for extraction, it was not useful.

This method can determine the peroxide content of samples, but theoretically there should be none left after the TCEP reduction process. Derivitization of the final product with FAME and

MSTFA analysis allowed us to quantitate the yield for the reactions. Since the FAME procedure took 24 hours before analysis, we decided to continue to use MSTFA since this method required only 30 minutes. MSTFA was also useful in determining the regioselectivity of linoleic acid oxidation (Figure 3-6). When analyzing the mass spectrum, we can determine the location of the hydroxyl group based on the fragmentation pattern. The desired product (13-(S)- hydroxyoctadeca-9Z,11E-dienoic acid) should show peaks at m/z 173 and 369 (Figure 3-6), while the silyl group will show a major peak at m/z 73.109

Figure 3-6. Mass spectrum analysis of 13-(S)-hydroxyoctadeca-9Z, 11E-dienoic acid

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According to the GC/MS data, the expected product was made by the lipoxygenase- catalyzed oxidation of linoleic acid and that the regiochemistry corresponds to the C13 position.

GC/MS analysis of MSTFA-derived products was used to monitor all subsequent reactions in this study.

Optimization of the Lipoxygenase Reaction with Linoleic Acid

Effects of the type of lipoxygenase used in the biotransformations

After initial trials showed that the commercial lipoxygenase enzyme successfully oxidized low concentrations of linoleic acid, optimization trials were conducted to increase the volumetric productivity of the process. Because commercially-obtained enzyme would be cost- prohibitive for scale-up, we isolated a semi-purified form of the enzyme directly from soybeans.

Some was saved as ammonium sulfate suspensions, while the rest was lyophilized and stored at

-20°C. The ammonium sulfate suspensions had slightly increased activity compared to the commercially available lipoxygenase (Figure 3-7). Interestingly, the lyophilized preparation was even more active than the ammonium sulfate suspensions. In addition, it was more stable.

Figure 3-7. Comparison of lipoxygenase conversion of 1 mM linoleic acid

When increasing the initial concentration of linoleic acid to 5 mM, the isolated lipoxygenase produced 1.18 mM of product (23% yield) while the lyophilized preparation

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produced 0.83 mM product (17% yield). Adding a second portion of enzyme resulted in a 20% increase in linoleic acid conversion. All subsequent reactions therefore used this double addition strategy. After evaluating various lipoxygenase concentrations, a total of 8 mg of enzyme (in 2 separate additions) were optimal for a 50 mL reaction mixture. The optimal amount of lipoxygenase for the biotransformation of linoleic acid was determined to be 0.16 μg/mL of the lyophilized enzyme.

Effects of TCEP in the reaction mixtures

We found that when TCEP was included in the mixture throughout the reaction, the conversion rate increased compared to trials when it was added after the reaction was complete.

Starting with 5 mM linoleic acid, the isolated lipoxygenase produced 1.5 mM product (30% yield), which is only slightly higher than the conversion observed in the absence of TCEP. When lyophilized lipoxygenase was used, 2.5 mM product was observed (50% yield). This was a significant increase over the reactions without included TCEP, which was 0.83 mM, 17% yield

(Figure 3-8). It was ultimately determined that to reach maximum conversion with these reactions, an equimolar quantity of TCEP relative to the linoleic acid was required. This was expected since TCEP is a reducing agent for the hydroperoxide group of the product. In all subsequent reactions, TCEP was included in the mixture from the beginning of the reaction.

Figure 3-8. Comparison of reactions with TCEP included and those without

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Effects of increasing the ethanol concentration in the reaction mixtures

The effect of varying EtOH concentrations was evaluated at a fixed concentration of 4 mM linoleic acid. Two stock solutions of linoleic acid (1 M and 0.5 M) were prepared for these studies. Using the 0.5 M stock solution increased the EtOH concentration by 50%. It was determined that doubling the EtOH concentration increased the conversion of linoleic acid in the reaction by roughly 24%. Reactions that used the 1 M stock of linoleic acid produced 2.84 mM

(71% yield) of the hydroxylated product, while the one that used the 0.5 M stock resulted in 3.81 mM (95% yield). It is believed that the additional EtOH aided in linoleic acid solubility in the reactions, but when higher concentrations of EtOH were evaluated the conversion rates decreased. This may be due to deleterious effects of EtOH on the enzyme. These conditions afforded the highest product concentration we were able to reach using our reaction system. The optimal reaction conditions were determined to be a 1:1 ratio of TCEP: linoleic acid, using a double addition of lipoxygenase, and starting with a 0.5 M stock solution of linoleic acid in

EtOH.

Other Substrates Tested for Lipoxygenase Activity

Our industrial partner isolated their own linoleic acid samples and donated 250 mL to our laboratory. In addition, they donated a sample of orange seed oil, which has high concentrations of the triglyceride form of linoleic acid (45%). Each of these was tested for their ability to be accepted by the lipoxygenase enzyme. The isolated linoleic acid given to us by our partner had the same conversion rates as the purified linoleic acid purchased through Sigma. Interestingly, the triglyceride form of linoleic acid was also accepted by lipoxygenase. A 50% solution of the oil was prepared in ethanol. A 100 μL sample was added to a 50 mL reaction flask and resulted in a 40% conversion into the hydroxylated product. This was interesting because it showed that the enzyme also accepted a linoleic acid derivative lacking a free carboxylate.

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Thermal Cracking of 13S-Hydroxyoctadeca-9Z,11E-dienoic Acid to n-Hexanal

Once the lipoxygenase reactions had been optimized, we carried out pyrolysis studies of the product in an attempt to convert it to n-hexanal. This procedure closely resembled that used previously for castor oil pyrolysis. Because only small quantities of 13-hydroxy linoleic acid could be obtained from the lipoxygenase reactions, a heating medium was required for cracking purposes. Peanut oil was chosen because it had the highest smoking point of the oils evaluated

(230°C).110 We hoped that the n-hexanal would collect in a condenser after pyrolysis of the 13- hydroxy linoleic acid.

The lipoxygenase reaction residue isolated from a 500 mL reaction (1 g 13-hydroxy linoleic acid) was added to a microdistialltion flask and dissolved in 1 mL peanut oil for thermal cracking. Several products were observed in the GC chromatogram (Figure 3-9). The chromatogram obtained by heating pure peanut oil under the same conditions was subtracted from the reaction including 13-hydroxylated linoleic acid so that products derived from the fatty acids could be identified more easily.

Figure 3-9. GC difference chromatogram of the cracking products from lipoxygenase residue in peanut oil A) n-Hexanal, B) 2-pentyl furan, C) n-Nonanal and D) Lauric Acid

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We found that pyrolysis of the lipoxygenase product resulted in 0.16 mg of n-hexanal

(0.04% yield). Heating peanut oil alone (4 mL) resulted in 0.021 mg of n-hexanal. This was a significant difference, demonstrating clearly that cracking 13-(S)-hydroxyoctadeca-9Z, 11E- dienoic acid results in n-hexanal production. Since the lipoxygenase product mixture also included linoleic acid, we evaluated the cracking of this material alone (no peanut oil included).

It resulted in 0.14 mg of n-hexanal (0.09% yield). Since there was much more linoleic acid present in this control reaction as compared to the one with the lipoxygenase product, we concluded that only a small fraction of the n-hexanal was derived directly from linoleic acid.

Since free linoleic acid present in the pyrolysis starting mixture did not negatively affect n- hexanal production, no effort was put into separating it from hydroxylated linoleic acid. It is even possible that free linoleic acid may supplement the level of n-hexanal production from the pyrolysis reactions, although further studies would be required.

The major peak in the thermal cracking product mixture results from a rearrangement of

13-hydroxylinoleic acid (2-pentyl furan). This product was not observed when linoleic acid itself was heated. It is thought that having both alkenes remaining in the carbon backbone of the hydroxylated product leads to this rearrangement. This is further confirmed by the work of Min et al. who reported that 2-pentylfuran can be formed from linoleic acid through singlet and triplet oxygen species at increased pressure and temperatures.111 We hypothesize that a majority of the hydroxylated product is diverted by this rearrangement and was therefore not available to make n-hexanal.

It is worth mentioning that peanut oil produces large amounts of n-nonanal in its cracking products. Peanut oil is mostly composed of oleic acid, which when heated seems to break down into n-nonanal. This product was not observed in the lipoxygenase product cracking mixture or

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with linoleic acid alone. Ultimately, it would be much more desirable to carry out thermal cracking with no added carrier oil, which would yield a cleaner product mixture.

Conclusion

While literature states that yields of up to 80% and ~30 g 13-hydroperoxy linoleic acid can be reached with the lipoxygenase enzyme, we saw maximum yields of 90% (1.26 g) 13- hydroxy linoleic acid. A large part of this was due to the fact that a pressurized container is necessary to reach higher product titer values. We currently do not have access to this type of container and it would also need to have oxygen continuously added. The lipoxygenase product contains a peroxide and having that functional group under pressure can easily result in undesired side reactions. While TCEP was included in these reactions to reduce the peroxide in situ, there is always the possibility that some level would still remain. Unfortunately, a product titer of 4.5 mM (1.26 g) is too low to provide enough 13-hydroxylinoleic acid for a carrier-free pyrolysis. The requirement for an equimolar level of TCEP is also undesirable since this reagents falls outside the realm of acceptable reductants for “natural” flavor production. Glutathione peroxidase would have to be substituted, which would create a two-enzyme system, which is also undesirable.

The lipoxygenase route was the second strategy evaluated for converting linoleic acid into n-hexanal. While some n-hexanal was produced, its yield was depressed by the formation of a rearrangement product (2-pentyl furan) that cannot be converted into n-hexanal. The maximum amount of n-hexanal made from 1.26 g 13-hydroxy linoleic acid was 0.16 mg (0.04% yield).

While this was a promising result, it was not clear how the approach could be scaled up and optimized further. One key requirement identified here was to remove the second alkene to avoid forming the furan rearrangement product during the pyrolysis step.

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CHAPTER 4

PRODUCTION OF 13S-HYDROXY-9(Z)-OCTADECENOIC ACID USING LINEOLATE 13- HYDRATASE FROM L. ACIDOPHILUS

Background

The previous chapters have described two different strategies that we explored to convert linoleic acid to n-hexanal. The approach described here is similar to that described in the previous chapter; however, the enzyme chosen here is a hydratase, rather than a lipoxygenase.

The lipoxygenase-catalyzed conversion of linoleic acid leaves both alkenes present in the carbon backbone. By contrast, lineolate 13-hydratase adds water across the C12-C13 alkene, which adds a hydroxyl group at C13 but also eliminates one of the two unsaturations. This product more closely resembles that of ricinoleic acid found in castor oil. We hypothesized that the product of the C12-C13 hydratase reactions would therefore undergo cleaner and more efficient thermal cracking to n-hexanal. These three structures are compared in Figure 4-1. This chapter will explore our work with lineolate 13-hydratase and was our final attempt at converting linoleic acid to n-hexanal.

A) B)

C)

Figure 4-1. Comparison of the cracking substrates evaluated A) from castor oil, B) lipoxygenase and C) lineolate 13-hydratase

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Experimental Procedures

Cloning of pEA4

The L. Acidophilus strain KCTC 3164 lineolate 13-hydratase gene (Accession number:

KJ560553) was synthesized by GenScript and sent to us in a pUC57 plasmid vector. The pET15b plasmid was purified by a previous member of our group, which includes a His tag. Both plasmids were prepared on large scales and purified by CsCl density gradient ultracentrifugation in the presence of ethidium bromide. The hydratase gene was inserted into the pET-15b vector using NdeI and BamHI restriction enzymes. All linearized products were purified by agarose gel electrophoresis prior to ligation using a 3:1 (vector:insert) ratio. After ligation, the mixture was used to transform E. coli E10Blue cells. Plasmid DNA was isolated from randomly-chosen colonies and sequenced to verify the desired construct, which is designated pEA4. Plasmid pEA4 was used to transform E. coli T7 Express and BL21(DE3)-Gold cells.

Cloning of pEA6

The lineolate 13-hydratase gene was isolated from the previously mentioned pEA4 plasmid using AatII and NdeI restriction sites. The pACYC Duet-1 plasmid was a generous donation by the Butcher lab (University of Florida). Both plasmids were prepared on large scales and purified by CsCl density gradient ultracentrifugation in the presence of ethidium bromide.

The hydratase gene was inserted into the pACYCDuet-1 vector using AatII and NdeI restriction enzymes. All linearized products were purified by agarose gel electrophoresis prior to ligation using a 3:1 (vector:insert) ratio. After ligation, the mixture was used to transform E. coli

E10Blue cells. Plasmid DNA was isolated from randomly-chosen colonies and sequenced to verify the desired construct, which is designated pEA6. Plasmid pEA6 was used to transform E. coli T7 Express and BL21(DE3)-Gold cells.

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Lineolate 13-Hydratase Expression

Colonies of T7 Express cells carrying pEA4 were grown on LB AmpR plates and then a single colony was used to make a starter culture at 37ºC (10 mL of SOC containing 0.1 mg/mL ampicillin). A culture was then prepared by taking 10 mL of the overnight culture and adding it to 1 L of sterile SOC containing 0.1 mg/mL ampicillin. This culture was grown at 37°C and 250 rpm until an OD600~0.4 was reached, then it was placed into a 16°C shaker. Once an OD600~0.6 was reached, the culture was induced with 0.1mM IPTG. Once induced, the culture was incubated at 16°C for 18 hours at 250 rpm. The cells were collected by centrifugation at 6,000 rpm for 15 minutes at 4°C. They were washed twice with 20 mL 0.2 M NaCl and then resuspended in an equal amount (1 mL/g) of citrate-phosphate buffer pH 6. The cells were lysed using sonication and centrifuged at 16,000 rpm for 45 minutes at 4°C. The same expression, harvesting and lysis procedure was followed for an overexpression host carrying the pEA6 plasmid, except that chloramphenicol (35 μg/mL) was the antibiotic throughout.

Bicinchoninic Acid Assay

The manufacturer’s (Sigma) protocol was followed.

Lineolate 13-Hydratase Enzymatic Reactions

A 0.5 M linoleic acid stock was prepared in EtOH. The conversion of linoleic acid into

13-hydroxy-9Z-octadecenoic acid was performed in 50 mM citrate-phosphate buffer (pH 6) containing 50 mM linoleic acid. In a flask, 6 mL of the reaction mixture was incubated at room temperature for 24 hours with agitation at 300 rpm. For the biotransformation, 25 g/L whole cells or 1.0 mg/mL of crude cell lysate were used in each reaction. The reactions were subsequently extracted with EtOAc and evaluated via MSTFA derivitization and GC/MS analysis using

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EA1_METH. For larger scale reactions, a continuous extraction was necessary prior to extraction with EtOAc due to the formation of emulsions.

13-Hydroxy-9Z-Octadecenoic Acid Cracking Procedure

To a microdistillation flask, the lineolate 13-hydratase product residue was added along with 2 mL of peanut oil. Finally, a small amount of rosin was added to the flask. The condenser

(with water chilled to 4°C) was added to the flask and the resulting mixture was submerged into a crucible filled with sand. Sand was used to reach higher temperatures, since peanut oil would begin to smoke around 220°C.110 A thermometer was placed in the sand bath, which was heated by a Bunsen burner. The temperature was slowly raised to 250°C, which is when the residue bubbling inside the flask ceased and considerable smoking occurred. The oil was allowed to cool and the condenser was washed with EtOAc. The EtOAc portions were combined and 1 mM lauric acid was added. The sample was added to a crimp top vial for GC/MS analysis via

JON_METH. For larger scale reactions, no peanut oil was included in the cracking medium.

GC/MS analysis: EA1_METH

GC/MS was performed on a Hewlett-Packard (HP) 5890 Series II Plus gas chromatograph (He carrier gas; 1.0 mL/min; split/splitless injector 220ºC; injection volume 1µL) with a DB-17 column (30m long; 250 µm i.d; 0.25 μm thickness). The initial temperature was set to 60°C for 2 minutes (3.5 minute solvent delay). The temperature was programmed from 60ºC at 10ºC/minute to 250ºC (hold for 10 minutes). The instrument was equipped with an auto- sampler. The GC was coupled to a HP 5971 series mass selective detector.

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Results and Discussion

Cloning and Expression of Lineolate 13-Hydratase

Park et al. previously reported that the pACYC-Duet1 plasmid vector was optimal for the expression of lineolate 13-hydratase.77 They determined that the pACYCDuet-1 plasmid resulted in a 1.6 fold increase in enzymatic activity over the pET-15b plasmid. The pET-15b plasmid is commonly used in our laboratory and was previously purified by another member of the group, while pACYC-Duet 1 was originally not easily accessible. pET-15b was therefore chosen as the initial expression plasmid vector for lineolate 13-hydratase due to its frequent use in our laboratory. In addition, a 1.6 fold difference in enzymatic activity was determined to be close enough for initial studies.

Figure 4-2. Construction of the pEA4 plasmid from pET-15b and pUC57

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The lineolate 13-hydratase gene (1782 base pairs) was synthesized by GenScript and sent to us in a pUC57 plasmid vector. The synthesis of the pEA4 plasmid is described in Figure 4-2.

The pEA4 plasmid overexpresses the hydratase protein with an N-terminal His tag fusion, which can be easily removed through its thrombin cleavage site. The fusion tag is useful for purification purposes since the procedure is straight forward. Lineolate 13-hydratase was observed at 69.6 kDa, which is slightly increased over the native size (68 kDa) due to the His tag and thrombin cleavage site.77

Initial overexpression studies used LB medium and 30°C along with BL21(DE3)-Gold cells as the host. This is the standard overexpression approach used by our lab. It was discovered that a majority of the protein was expressed as inclusion bodies (misfolded protein). Most proteins are not active in their inclusion body form, including lineolate 13-hydratase. We therefore attempted eliminate these inclusion bodies and obtain soluble protein. Park et al. undertook a similar study and found that SOC medium (40 mM Mg2+), E. coli ER2566 cells, expression at 16°C, and an 18 hour induction time was optimal for this enzyme.77 This was the starting point for our investigation. The results of our study supported those previously reported.77 Therefore, for further studies, plasmids encoding lineolate 13-hydratase were used to transform T7 Express E. coli cells (New England Biolabs) and grown in SOC medium, then induced at 16°C for 18 hours. The difference in overexpression systems can be observed in

Figure 4-3. It is clear that the T7 Express cells lead to an increase in protein expression; however, there are still a large amount of inclusion bodies present. Since there is a significant amount of soluble protein (Figure 4-3), the inclusion bodies were overlooked.

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Figure 4-3. Comparison of the soluble protein present in pEA4 overexpression systems a) T7 Express and b) BL21(DE3)-Gold cell lines

It was later determined that the cells needed to be permeabilized before use, otherwise a decrease in activity of the lineolate 13-hydratase was observed. Kang et al. had also come to this conclusion in 2016 and found that the best permeabilzer for this purpose was 0.2 M NaCl.76

Once expressed, the E. coli cells were washed twice with 0.2 M NaCl prior to use in linoleic acid hydration reactions or prior to lysing.

While performing reactions, it was discovered that the T7 express cells transformed with pEA4 died quickly and could not be stored as glycerol stocks at -80°C. This led to a continuous decrease in lineolate 13-hydratase overexpression with complete loss of overexpression after 2 weeks. Ultimately, this led to the synthesis of plasmid pEA6 which was designed to be more stable in the T7 express cells.77 The lineolate 13-hydratase gene (2127 base pairs) was therefore subcloned from pEA4 using NdeI and AatII (Figure 4-2). The pACYC Duet-1 plasmid carries a pET-15b replicon and is designed for the coexpression of two target genes. The lineolate 13- hydratase gene was inserted after the second T7 promoter, thereby keeping both T7 promotors in the plasmid sequence. Hydratase overexpression by the pEA6 plasmid can be observed in Figure

4-4. The resulting expressed lineolate 13-hydratase was 67.4 kDa, which is within experimental error of the 68 kDa reported in literature.77

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Figure 4-4. Construction of the pEA6 plasmid from pACYC Duet-1 and pEA4

Test inductions were performed with pEA6 at 16°C, 25°C, and 37°C in the same manner as described for the strains harboring the pEA4 plasmid (Figure 4-5). The optimum induction conditions were found to be 16°C, SOC medium and 18 hours which are consistent with those reported by Park et al.77 In addition, it was discovered that the amount of inclusion bodies formed was greatly decreased when compared to strains carrying the pEA4 plasmid.

Figure 4-5. Soluble proteins present in pEA6 18 hour overexpression A) 16°C B) 25°C and C) 37°C

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Detection of 13S-Hydroxy-9Z-Octadecenoic Acid

The products from the reactions catalyzed by lineolate 13-hydratase were derivitized with

MSTFA and analyzed via GC/MS. By analyzing the mass spectrum of the products, we could determine the regiochemistry of hydration based on the fragmentation pattern. The expected 13- hydroxy-9Z-octadecenoic acid product will show peaks at m/z 173 and 371 (Figure 4-6). The m/z 371 species undergoes further fragmentation, making it less abundant. Our spectral data match those reported by Oh et al. for 13-hydroxy-9Z-octadecenoic acid.84 The large peak at m/z

109 73 is due to the silyl group. Based on these data, we concluded that C13 bears the hydroxyl group.

Figure 4-6. Mass spectrum analysis of 13-hydroxy-9Z-octadecenoic acid

Hydration of Linoleic Acid with Crude Lysate Suspensions

The concentration of total protein in the crude lysate was measured by a BCA assay. In order to determine the optimal concentration of crude lysate for preparative reactions, it was added between 0 and 2 mg/mL and the production of 13-hydroxy-9Z-octadecenoic acid from

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linoleic acid was determined (Figure 4-7). These reactions were carried out at pH 6.0, 25°C, 300 rpm shaking and 50 mM linoleic acid (from a 1 M stock dissolved in EtOH). The reactions were monitored at 6 and 24 hours for reaction progress.

It was determined that the optimal level of total protein from crude lysate for hydrating

50 mM linoleic acid into 13-hydroxy-9Z-octadecenoic acid was 1 mg/mL. At higher levels, the conversion leveled off at around 76% for reasons that were not explored.

Figure 4-7. Determination of the optimal crude lysate concentration for hydratase reactions

In parallel, negative controls were carried out with a crude extract from T7 Express E. coli cells lacking a plasmid to ensure there was no linoleic acid hydration without the overexpressed enzyme. In addition, reactions were also carried out without any added lysate

(containing only linoleic acid and buffer). All negative controls showed no detectable linoleic acid hydration. We therefore concluded that lineolate 13-hydratase was required for producing

13-hydroxy-9Z-octadecenoic acid.

In order to determine the optimum linoleic acid concentration for the hydration reactions, starting concentrations ranging from 8 to 100 mM were evaluated. The reactions were monitored at both 6 and 24 hours and both single and double crude lysate additions were explored (Figure

4-8). When added, the second portion of crude lysate was introduced after 6 hours.

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Based on these results, we determined that the conversion rate of linoleic acid into 13- hydroxy-9Z-octadecenoic acid was linear up to 60 mM, where it began to level off when two portions of crude lysate were added. It is unknown why this behavior occurs, but substrate saturation of the enzyme is one possibility.77 It was clear that two enzyme additions were necessary to reach the maximum linoleic acid conversions. It was decided to run subsequent crude lysate reactions at 50 mM linoleic acid, because this showed the most reproducible conversion rates.

Figure 4-8. Determination of optimal linoleic acid concentation for crude lysate reactions

Once the optimized reaction conditions were developed, crude lysates from E. coli strains harboring pEA4 and pEA6 were compared. The crude lysate reaction from the former resulted in

73% conversion while the later afforded 62% conversion. Since the pEA4 plasmid yielded consistently better results, subsequent reactions using crude lysates relies on a strain carrying this plasmid.

Hydration of Linoleic Acid with Whole Cells

Whole cell biotransformations are preferable for reactions with toxic or hydrophobic substrates and this strategy is commonly utilized for reactions that oxygenate fatty acids, carboxylic acids, and aliphatic alkanes.77 The whole cell reactions performed in this study were

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carried out using 50 mM citrate-phosphate buffer, pH 6.0 at 25°C, 50 mM linoleic acid, 25 g/L cells and 0.25% (v/v) Tween 20. This concentration of cell mass was chosen due to a study by

Park et al. who reported that 25 g/L was the optimum cell concentration for hydration of linoleic acid.77 When using the whole cells that express the lineolate 13-hydratase from pEA4, a relatively low conversion of linoleic acid was observed (50 to 60%). It was later determined that the cells quickly lost activity during storage at -20°C. When using freshly permeabilzed cells, the conversion of linoleic acid reached a maximum of 75% in 48 hours; however, the same cell stock became inactive after 1 week of storage at -20°C.

When using freshly permeabilized cells that express the lineolate 13-hydratase from pEA6, the conversion of linoleic acid reached 76% in less than 24 hours. This showed that pEA6 led to a higher activity in T7 Express cells than pEA4. This result is further supported by the reports of Park et al.77 Due to the shorter reaction time, T7 Express cells harboring pEA6 were employed for all subsequent whole cell-mediated reactions. A key advantage of whole cell reactions is that they are almost infinitely scalable. Furthermore, such reactions are also more convenient because no lysis step is required.

Figure 4-9. Determination of the optimal linoleic acid concentration for whole cell reactions

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The optimum concentration of linoleic acid for whole cell reactions was explored using whole cells of T7 Express cells containing the pEA6 plasmid. As seen in Figure 4-9, the optimum linoleic acid concentration was 250 mM. Above 250 mM, the conversion consistently decreased. While the 13-hydroxy-9Z-octadecenoic acid concentration never actually leveled off, the conversion was not high enough to justify reactions with initial linoleic acid concentrations higher than 250 mM. As observed with the crude lysate reaction data, the maximum conversion of linoleic acid into 13-hydroxy-9Z-octadecenoic acid is 80% and it is unknown as to why this value could never be exceeded. We decided to carry out small-scale reactions at 50 mM but larger-scale reactions at concentrations of linoleic acid up to 210 mM.

Various Buffer Systems in the Hydratase Reactions

Using the optimized conditions developed for reactions with crude lysates, large amounts of emulsions formed when the reaction mixtures were extracted with EtOAc. The use of volatile buffers that could be removed by lyophilization was explored as a strategy to avoid emulsions.

Both ammonium bicarbonate and ammonium acetate were tested. Once the reactions were complete, they were frozen at -80˚C and lyophilized overnight. For larger reaction volumes

(>100 mL), most water was initially removed by a high vacuum rotary evaporator prior to lyophilization. The residue containing the desired product was resuspended in EtOAc, filtered and the solvent was evaporated. The reactions containing ammonium bicarbonate buffer consistently yielded lower enzymatic activity when compared to the citrate-phosphate buffer used in the optimum conditions (Figure 4-10). On the other hand, reactions carried out in ammonium acetate buffer showed the same activity levels as the citrate-phosphate buffer.

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Figure 4-10. Buffer system comparison of 10 mM linoleic acid conversion

Unfortunately, the use of volatile buffers only provided 30 to 50% product recoveries.

The fate of the missing material is not clear; however, the low recoveries prompted us to abandon this approach and all subsequent reactions were performed in 50 mM citrate-phosphate buffer at pH 6.

Effects of Temperature on the Hydration of Linoleic Acid

Unless otherwise stated, the temperature study reactions were performed in 50 mM citrate-phosphate buffer, pH 6 along with 50 mM linoleic acid (from a 1 M stock in EtOH), 25 g/L whole cells, stirring at 300 rpm and either 25˚C or 37˚C. These whole cell reactions were prepared with lineolate 13-hydratase expressed from both the pEA4 in BL21(DE3)-Gold and T7

Express cell lines. The results of these reactions are shown in Table 4-1. Conversions using T7

Express cells were the most consistent and there was no observable difference in the conversion of linoleic acid between the two temperatures. While the BL21(DE3)-Gold transformants were more active at 37˚C protein overexpression by these cells is lower. Since room temperature was more convenient, whole cell-mediated reactions were carried out at 25˚C.

Table 4-1. Whole cell-mediated hydrations of linoleic acid at two temperatures

E. coli cells 25˚C 37˚C T7 Express 69% conversion 68% conversion BL21-gold (DE3) 48% conversion 72% conversion

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Reactions using crude lysate prepared from T7 Express host cells overexpressing lineolate 13-hydratase from pEA4 were also performed at 25˚C and 37˚C with single and double lysate additions (Table 4-2). Once again, it was determined that there was little difference between the two temperatures for the double addition of crude lysate. The single addition crude lysate reactions, however, were not as active and it is believed to be due to enzyme inactivation during the reaction time.

Table 4-2. Crude lysate-mediated hydrations of linoleic acid at two temperatures

Temperature Single addition Double addition 25˚C 66% conversion 77% conversion 37˚C 54% conversion 70% conversion

Finally, whole cell-mediated reactions that utilized T7 Express cells transformed with plasmid pEA6 were carried out. The reactions were evaluated using the previously-optimized reaction conditions (25 g/L cells) at both 25˚C and 37˚C. There was little difference in conversion after 6 hours when 50 mM linoleic acid was used (Table 4-3). The same results were observed at increased concentrations of linoleic acid. These results contradict those reported by

Park et al.77 Based on these results, we selected 25˚C as the reaction temperature since it was more convenient.

Table 4-3. Six hour whole cell-mediated hydrations of linoleic acid at two temperatures

Temperature 50 mM linoleic acid 150 mM linoleic acid 250 mM linoleic acid 25˚C 78% conversion 69% conversion 46% conversion 37˚C 81% conversion 71% conversion 49% conversion

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Multiple Enzyme Additions to the Hydratase Reactions

As mentioned previously, multiple enzyme additions were necessary to reach the maximum conversion of linoleic acid using crude lysates. In an attempt to obtain full conversion of linoleic acid, the number of crude lysate additions was varied to determine the effect on linoleic acid conversion at a substrate concentration of 50 mM (Figure 4-11). Each crude lysate aliquot addition was made after 6 hour increments.

Figure 4-11. Multiple crude lysate additions to the hydratase reaction

In each reaction, the conversion leveled off after 32 hours. This was determined to be the optimum reaction time for crude lysate reactions from the T7 Express cells transformed with plasmid pEA4. Two additions of crude lysate containing lineolate 13-hydratase proved to be the best strategy for these biotransformations. A single addition did not reach the same conversion while reactions employing three and four additions showed nearly the same conversion values as that using a single addition. The reason for this phenomenon remains unknown; however, all subsequent hydrations of linoleic acid using crude lysates utilized a double addition strategy.

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Time Course Study for the Hydratase Reactions

Unless otherwise stated, the time course study reactions were performed at 25°C in 50 mM citrate-phosphate buffer, pH 6.0, 50 mM linoleic acid, 0.25% (v/v) Tween 20, and either

0.75 mg/mL crude lysate or 25 g/L whole cells. The reactions were evaluated for a 48 hour time frame in each case (Figure 4-12).

Figure 4-12. Time course study for the hydration of 50 mM linoleic acid

In Figure 4-12, it is shown that the most efficient reaction is the whole cell biotransformation from host cells transformed with pEA6. Within 6 hours, the conversion leveled out at 76% and remained at this level until the experiment was terminated. This was significant because it showed that the product did not decompose during extended reaction times.

In addition, this reaction also showed the highest conversion of those evaluated. The crude lysate reaction from cells transformed with the pEA4 plasmid reached its maximum conversion (72%) after 24 hours before leveling off. The other reactions reached their maximum conversions after

32 hours prior to leveling off. The crude lysate reaction from cells transformed with pEA6 plasmid had the worst conversion (68%), and the level actually declined after this level was reached. Based on these results, all subsequent reactions used whole cells of T7 Express transformed with pEA6 6-8 hours at 25˚C.

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Addition of Detergent in the Reaction Mixtures

Initially, reactions were run with no detergent and the linoleic acid was dissolved in ethanol, prior to its addition to the aqueous reaction medium. It was ultimately determined that dissolving the substrate in ethanol had no advantages in the reaction system, which was also supported by the results of Park et al.77 When detergent was added to the reactions, the hydration of linoleic acid increased. Lower detergent concentrations (ca. 1%) resulted in higher conversions (67%) than when higher detergent concentrations were employed (50%). This was consistent with results reported by Park et al.77 Furthermore, adding detergent to the reaction mixtures, the extraction procedures also decreased emulsion formation. For this reason, 0.25%

(v/v) Tween 20 was used in the bioconversions.

Extraction Methods to Remove Biotransformation Products

When either the crude lysate or whole cell-mediated reactions were extracted with

EtOAc, large amounts of emulsion formed. Such emulsions typically form when doing oil in water extractions and techniques for breaking up emulsions include adding salt or detergents or centrifugation. While the latter is preferred, most labs do not have the equipment for large scale centrifugations involving flammable mixtures.112 We explored several approaches to avoid forming emulsions in our reaction mixture: continuous extraction (with and without added heat), charcoal adsorption, centrifugation and simple solvent extraction.

The first method evaluated was a traditional continuous extractor that used heat to continuously distill fresh EtOAc into the reaction mixture (Figure 4-13). Three days and several additions of fresh EtOAc were required to extract the lineolate 13-hydratase reaction product completely from the reaction mixture. The round bottom flask containing EtOAc was constantly under heat (~100°C to evaporate EtOAc) and this degraded the hydratase product into a brown

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residue. This method was abandoned because of the long time required and the product decomposition observed.

Figure 4-13. Continuous extraction apparatus using heat113

Centrifuging the mixture obtained by shaking with EtOAc efficiently extracted 13- hydroxy-9Z-octadecenoic acid on a small scale; however, once the reactions were scaled up

(more than 100 mL), the centrifugation process became tedious and time consuming. In principle, this could be overcome with a larger explosion-proof centrifuge; however, because the required equipment was not available in our lab, this approach was not explored further.

Charcoal was tested as a way to remove 13-hydroxy-9Z-octadecenoic acid directly from the reaction mixture. Cason and Gillies studied the adsorption of fatty acids onto charcoal.114

They found that linoleic acid was adsorbed by charcoal and could be eluted with 95% EtOH.

When we attempted to replicate this procedure, even after 3 extractions and washing the charcoal with 95% EtOH, linoleic acid was still present in the aqueous reaction mixture. Since EtOH was present in the reaction mixture, we repeated the charcoal adsorption without added EtOH. While a slight increase in linoleic acid adsorption was observed, a significant amount still remained in

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the aqueous mixture. Due to these difficulties, this approach was discontinued although we did find that charcoal adsorbed the linoleic acid hydration product with low efficiency.

Finally, a continuous extraction method that did not require heating was tested for product removal from the reaction mixture. Ethyl acetate was recirculated by an HPLC pump into a flask containing the aqueous reaction mixture. After bubbling upward, the overflow was directed back into the HPLC solvent reservoir (Figure 4-14).

Figure 4-14. Continuous extraction set up using an HPLC pump and EtOAc, photo by author

As the extraction proceeded, a clear separation of layers was observed, including a layer of emulsion (Figure 4-15). The aqueous portion could then be removed and easily extracted using a separatory funnel. The continuous extraction using the HPLC pump was the most efficient method tested for lineolate 13-hydratase product removal. It is easily scalable to any foreseeable volume and we were able to consistently extract 90% of the desired product.

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Figure 4-15. Continuous extraction layer separation with an HPLC pump, photo by author

Cracking of Biotransformation Products from Lineolate 13-Hydratase Reactions

To determine whether n-hexanal could survive our proposed thermal cracking procedure, n-hexanal was dissolved in 3 mL of linoleic acid to a final concentration of 5 mM. A small amount of rosin was added and the apparatus described previously (Figure 3-3) was placed in a sand bath and heated by a bare flame. After the cracking was complete, the condenser was washed with EtOAc. Lauric acid was added as an internal standard to the combined EtOAc fractions at a final concentration of 1 mM, then the sample was analyzed by GC/MS. It was observed that all of the n-hexanal was recovered and that n-hexanal formed during the thermal cracking process would be stable to the reaction conditions.

The first attempt at applying this procedure to the lineolate 13-hydratase product used peanut oil as a heating medium because only small quantities of the product were then available.

To monitor the cracking temperature, peanut oil was also placed into a crystallization dish with a thermometer. Later, a sand bath was substituted due to the smoke that occurred from the peanut oil. These yielded 0.48-0.80 mM n-hexanal from 0.71 g 13-hydroxy linoleic acid, which corresponded to 0.060-0.106 mg (0.02-0.04% yields) of pure n-hexanal. The chromatogram for the cracking products was cluttered due to products from both the hydratase reaction as well as peanut oil that had collected on the condenser (Figure 4-16).

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Figure 4-16. Chromatogram of cracking products dissolved in peanut oil

In Figure 4-16, the main product from 13-hydroxy-9Z-octadecenoic acid cracking is indicated (n-hexanal) at 2.50 minutes. The second product indicated is the main cracking product from peanut oil (n-nonanal) at 6.80 minutes. This was determined by subtracting the peanut oil chromatogram from that of the hydratase reaction mixture with peanut oil. When the cracking procedure was performed on peanut oil, 0.14 mM n-hexanal (0.017 mg) was produced, which was much less than from the hydratase reaction product. This is strong evidence that the majority of n-hexanal is derived from the lineolate 13-hydratase product. In addition, when 3.6 g of linoleic acid was cracked alone (no peanut oil), 0.93 mM n-hexanal (0.116 mg, 0.01% yield) was produced. It was determined that there was no need to separate linoleic acid from the 13- hydroxy-9Z-octadecenoic acid since the former had no negative impacts on n- hexanal production. Finally, when 13-hydroxy-9Z-octadecenoic acid was dissolved in peanut oil and cracked, the same n-hexanal titers were reached as when no peanut oil. We thus concluded that

13-hydroxy-9Z-octadecenoic acid was the dominant contributor to n-hexanal production, as we had hoped.

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When lineolate 13-hydratase reactions were scaled up to 500 mL or more, the quantities of 13-hydroxy-9Z-octadecenoic acid were sufficiently high (1.58 g) that peanut oil was no longer needed as a heating medium in the cracking mixture. After adding rosin, heating the mixture, and washing the condenser with EtOAc, 1.38 mM n-hexanal (3.10 mg, 0.6% yield) was produced.

The resulting chromatogram was much cleaner than those obtained when peanut oil was present

(Figure 4-17).

Figure 4-17. Chromatogram of carrier-free hydratase product cracking

More n-hexanal was produced from the carrier-free cracking procedure as compared to reactions in which the starting material was dissolved in peanut oil. In Figure 4-17, it can be seen that the main product is n-hexanal at 2.50 minutes. In addition, at 4.80 minutes, a rearrangement product (2-pentyl furan) was formed at an equal concentration as n-hexanal. Finally, there is a product observed at 3.41 minutes, which is predicted to be α-pinene derived from heating rosin at high temperatures. It is worth mentioning that the n-nonanal peak was not present at 9.0 minutes, confirming that this substance is derived from peanut oil. When undergoing the cracking procedure, it was observed that the liquid that collected in the condenser cup was also getting heated and re-condensing (Figure 3-3). n-Hexanal is known to be a reactive and volatile aldehyde; it is therefore possible that its being constantly heated would cause it to react further. It

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is believed that higher n-hexanal titers might be achievable if the n-hexanal product were protected from further heating.

Once the linoleic acid hydratase reactions had been scaled up at high substrate concentrations, sufficient product was available to carry out the cracking procedure in a 3-neck flask, thereby eliminating the possibility of reheating the n-hexanal product (Figure 4-18).

Figure 4-18. 3-neck flask cracking apparatus105

A bare flame was used to heat the hydratase product mixed with rosin in the 3-neck flask.

The collection flask was placed in an ice bath to ensure that any volatile products collected did not react further or evaporate. The resulting chromatogram was even cleaner than those obtained previously (Figure 4-17). Using 20 mL of the lineolate 13-hydratase reaction product for the cracking procedure (11 g 13-hydroxy-9Z-octadecenoic acid), 38.9 mg (1% yield) of n-hexanal was produced (Figure 4-19). The rearrangement product (2-pentyl furan) was also present at 4.80 minutes and α-pinene was present at 3.40 minutes. To the best of our knowledge, this is the first example of an all-natural process producing these high titers of n-hexanal.

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Figure 4-19. Chromatogram of 3-neck flask cracking products

Finally, a thermal cracking procedure was done on the hydratase product mixture (2 g 13- hydroxy-9Z-octadecenoic acid) without added rosin. This resulted in lower n-hexanal production

(1.94 mg, 0.3% yield) then when rosin was present. It was therefore determined that rosin aids more in thermal cracking than just making free fatty acids and is necessary to reach the higher n- hexanal quantities. At this time, the exact function of the rosin is unknown.

Scaling Up Lineolate 13-Hydratase Reactions

The optimum reaction conditions along with the extraction and cracking procedures have been developed in previous sections. For industrial purposes, these procedures need to be scaled up to liters; therefore, reactions containing a starting concentration of linoleic acid up to 210 mM were scaled up to 100 mL using whole cell catalysis. The conversion of the scaled up reactions remained at 80% and the extraction and cracking procedures were easily applied to these volumes. This shows that a significant amount of hydrated linoleic acid can be produced at a

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single time. At the present time, the need for hundreds of grams of linoleic acid is a practical barrier to further scale up.

Other Substrates Evaluated with Lineolate 13-Hydratase

Several substrates were evaluated with lineolate 13-hydratase including oleic, palmitoleic, linolenic, eladic, palmitic and lauric acids. These reactions were performed in 50 mM citrate-phosphate buffer, pH 6.0, 25˚C, 0.75 mg/mL crude lysate, and 10 mM fatty acid (1

M stocks in EtOH). No conversion was observed with any of the fatty acids evaluated with the exception of linoleic acid (80% conversion). These results are consistent with those reported by

Oh et al.84

The same series of substrates were also used in whole cell biotransformations in 50 mM citrate phosphate buffer, pH 6.0, 25˚C, 10 mM fatty acid, 0.25% (v/v) Tween 20 and 25 g/L whole cells. Unlike reactions using crude lysate containing lineolate 13-hydratase, a few acids exhibited small amounts of hydration including palmitoleic acid (16%), linolenic acid (25%), and methyl lineolate (14%) after 24 hours. The hydrated products were all confirmed by analyzing their MS data in a manner analogous to that used for linoleic acid (Appendix C). The remaining substrates showed no detectable hydration products at the limits of GC/MS detection. To the best of our knowledge, this is the first example of lineolate 13-hydratase accepting a substrate other than linoleic acid. With reaction optimization, it might be possible to develop these alternate substrates into useful processes; however, this was not investigated in our study.

Future Directions

The greatest remaining need is to optimize the cracking procedure. The maximum n- hexanal yield reached is 1% from our current process. This results in a significant amount of wasted hydrated linoleic acid; therefore, effort needs to be put into optimizing this procedure prior to further scaling up the reaction. Ultimately, the lineolate 13-hydratase whole cell

138

reactions need to be increased to the liter scale. The optimum reaction conditions have been developed along with an efficient extraction procedure and analytical methods. Scaling up to liters would allow for higher 13-hydroxy-9Z-octadecenoic acid quantities to be produced and this will allow cracking of larger volumes. This would allow a different apparatus that might give a more controlled pyrolysis process and better n-hexanal recovery.

Lineolate 13-hydratase has been found to be unstable under our storage conditions. One future direction would be to engineer this protein to increase its stability and increase the acceptance of other fatty acid substrates. By increasing the substrate scope, this enzyme could be used to hydrate a wider variety fatty acids to produce various aldehyde compounds via thermal cracking. Although there is no current crystal structure for this enzyme, a similar one (oleate hydratase) has been crystallized and this information might be applied to lineolate 13-hydratase due to their high sequence similarity. Crystallization of lineolate 13-hydratase would be of great use to further understand the reaction mechanism and substrate binding of this enzyme.

Conclusion

In this chapter, various reaction conditions were explored for lineolate 13-hydratase.

Optimum reaction conditions were developed (50 mM citrate-phosphate buffer, pH 6.0, 25˚C,

0.25% (v/v) Tween 20, 50 mM linoleic acid, and 25 g/L whole cells expressing lineolate 13- hydratase from pEA6 in E. coli T7 Express) along with efficient extraction procedures and analytical methods. We have now shown that measureable quantities of n-hexanal can be produced through our cracking procedures. n-Hexanal can also be further purified through fractional distillation due to its low boiling point (~69˚C); however, this has yet to be attempted in our lab. To the best of our knowledge, this is the first example of an all-natural, single-enzyme process that can produce n-hexanal from linoleic acid. To date, low concentrations of n-hexanal

(micromolar range) have been produced through the lipoxygenase and lyase method. This is the

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only other known natural method that has produced significant amounts of n-hexanal from linoleic acid.

While scaling this process up to liters has yet to be accomplished, the procedures are relatively simple so it should be achievable. In addition, whole cell catalysis was determined to be the most efficient at producing 13-hydroxy-9Z-octadecenoic acid, and this is very straightforward to scale up. The hydration product is essential to efficient n-hexanal production, since pyrolyzing free linoleic acid gives only traces of the desired product. Finally, we have also shown that thermal cracking of peanut oil gave large quantities of n-nonanal, which has not been reported previously to the best of our knowledge.

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APPENDIX A ADDITIONAL PLASMID INFORMATION

Nucleic Acid Sequences

CCCTTCGCTACAAGCCTACAAGTCATCTCGCCGCAACCGGAACCCGCAGCCATGGG GACGGAGGCGGAGCAGCCGGACATGGACAGCCACCGAAACGACGGCGTCGTGGTG GTGCCAGCGCCGCGCCCGCGTAAGGGGCTCGCTTCCTGGGCGCTCGACCTGCTTGAG TCCCTCGCCGTGCGCCTCGGCCACGACAAGACCAAGCCGCTCCACTGGCTCTCCGGC AACTTCGCCCCCGTCGTCGAGGAGACCCCGCCGGCCCCAAACCTTACCGTCCGCGGA CACCTCCCGGAGTGCTTGAATGGAGAGTTTGTCAGGGTTGGGCCTAATCCGAAGTTT GCTCCTGTTGCGGGGTATCACTGGTTTGATGGAGACGGGATGATTCATGCCATGCGT ATTAAGGATGGAAAAGCTACCTATGTATCAAGATATGTGAAGACTGCCCGCCTCAA ACAAGAGGAGTATTTTGGTGGAGCAAAGTTTATGAAGATTGGAGACCTTAAGGGAT TTTTTGGATTGTTTATGGTCCAAATGCAGCAACTTCGGAAAAAATTCAAAGTCTTGG ATTTTACCTATGGATTTGGGACAGCTAATACTGCACTTATATATCATCATGGTAAACT CATGGCCTTGTCAGAAGCAGATAAGCCATATGTTGTTAAGGTCCTTGAAGATGGAGA CTTGCAGACTCTTGGCTTGTTGGATTATGACAAAAGGTTGAAACATTCTTTTACTGCC CATCCAAAGGTTGACCCTTTTACAGATGAAATGTTCACATTCGGATATTCACATGAA CCTCCATACTGTACATACCGTGTGATTAACAAAGAAGGAGCTATGCTTGATCCTGTG CCAATAACAATACCGGAATCTGTAATGATGCATGATTTTGCCATCACAGAGAATTAC TCTATTTTTATGGACCTCCCTTTATTGTTCCGACCAAAGGAAATGGTGAAGAACGGT GAGTTTATCTACAAGTTTGATCCTACAAAGAAAGGTCGTTTTGGTATTCTCCCCCGCT ATGCAAAGGATGACAAACTCATCAGATGGTTTCAACTCCCTAATTGTTTCATATTCC ATAATGCTAATGCTTGGGAAGAGGGTGATGAAGTTGTTCTAATTACCTGCCGCCTTG AGAATCCAGATTTGGACAAGGTGAATGGATATCAAAGTGACAAGCTCGAAAACTTC GGGAATGAGCTGTACGAGATGAGATTCAACATGAAAACGGGTGCTGCTTCACAAAA GCAATTGTCTGTTTCTGCTGTGGATTTTCCTCGTGTTAATGAGAGCTATACTGGCAGA AAGCAGCGGTATGTCTACTGCACTATACTTGACAGCATTGCGAAGGTGACTGGCATC ATAAAGTTTGATCTGCATGCTGAACCGGAAAGTGGTGTGAAAGTACTTGAAGTGGG AGGAAATGTACAAGGCATATATGACCTGGGACCTGGTAGATTTGGTTCAGAGGCGA TTTTTGTTCCCAAGCATCCAGGTGTGTCTGGAGAAGAAGATGACGGCTATTTGATAT TCTTTGTACACGACGAGAACACAGGGAAATCTGAAGTAAATGTTATCGATGCAAAG ACAATGTCTGCTGATCCAGTTGCGGTGGTTGAGCTTCCTAATAGGGTTCCTTATGGA TTCCATGCCTTTTTTGTAACTGAGGACCAACTGGCTCGACAGGCGGAGGGGCAGTGA AGATACGGCACCTGCAGATTCTGCACACGCGGGTACAGGTTGGAAATTATTGCAGG GCATGTATATGTATGGGACAAGTTTATTACACATGTATTCGAACCACACATTACACA AGTTTATTGCAGGACGTGTATTCGAA Figure A-1. Sequence of Zea mays CCD1 (Accession: DQ100346)

CATATGGCGGAGAAACTCAGTGATGGCAGCGTCATCATCTCAGTCCATCCTAGACCC TCCAAGGGTTTCTCCTCGAAGCTTCTCGATCTTCTCGAAAGACTTGTCGTCAAGCTCA TGCACGATGCTTCTCTCCCTCTCCACTACCTCTCAGGCAACTTCGCTCCCATCCGTGA TGAAACTCCTCCCGTCAAGGATCTCCCCGTCCATGGATTTCTTCCCGAATGCTTGAAT GGTGAATTTGTGAGGGTTGGTCCAAACCCCAAGTTTGATGCTGTCGCTGGATATCAC TGGTTTGATGGAGATGGGATGATTCATGGGGTACGCATCAAAGATGGGAAAGCTAC Figure A-2. Sequence of Arabidopsis thaliana CCD1 (Accession: NM_116217)

141

(Figure A-2. continued) TTATGTTTCTCGATATGTTAAGACATCACGTCTTAAGCAGGAAGAGTTCTTCGGAGC TGCCAAATTCATGAAGATTGGTGACCTTAAGGGGTTTTTCGGATTGCTAATGGTCAA TATCCAACAGCTGAGAACGAAGCTCAAAATATTGGACAACACTTATGGAAATGGAA CTGCCAATACAGCACTCGTATATCACCATGGAAAACTTCTAGCATTACAGGAGGCAG ATAAGCCGTACGTCATCAAAGTTTTGGAAGATGGAGACCTGCAAACTCTTGGTATAA TAGATTATGACAAGAGATTGACCCACTCCTTCACTGCTCACCCAAAAGTTGACCCGG TTACGGGTGAAATGTTTACATTCGGCTATTCGCATACGCCACCTTATCTCACATACA GAGTTATCTCGAAAGATGGCATTATGCATGACCCAGTCCCAATTACTATATCAGAGC CTATCATGATGCATGATTTTGCTATTACTGAGACTTATGCAATCTTCATGGATCTTCC TATGCACTTCAGGCCAAAGGAAATGGTGAAAGAGAAGAAAATGATATACTCATTTG ATCCCACAAAAAAGGCTCGTTTTGGTGTTCTTCCGCGCTATGCCAAGGATGAACTTA TGATTAGATGGTTTGAGCTTCCCAACTGCTTTATTTTCCACAACGCCAATGCTTGGGA AGAAGAGGATGAAGTCGTCCTCATCACTTGTCGTCTTGAGAATCCAGATCTTGACAT GGTCAGTGGGAAAGTGAAAGAAAAACTCGAAAATTTTGGCAACGAACTGTACGAAA TGAGATTCAACATGAAAACGGGCTCAGCTTCTCAAAAAAAACTATCCGCATCTGCG GTTGATTTCCCCAGAATCAATGAGTGCTACACCGGAAAGAAACAGAGATACGTATA TGGAACAATTCTGGACAGTATCGCAAAGGTTACCGGAATCATCAAGTTTGATCTGCA TGCAGAAGCTGAGACAGGGAAAAGAATGCTGGAAGTAGGAGGTAATATCAAAGGA ATATATGACCTGGGAGAAGGCAGATATGGTTCAGAGGCTATCTATGTTCCGCGTGAG ACAGCAGAAGAAGACGACGGTTACTTGATATTCTTTGTTCATGATGAAAACACAGG GAAATCATGCGTGACTGTGATAGACGCAAAAACAATGTCGGCTGAACCGGTGGCAG TGGTGGAGCTGCCGCACAGGGTCCCATACGGCTTCCATGCCTTGTTTGTTACAGAGG AACAACTCCAGGAACAAACTCTTATATAACTCGAG Figure A-2. Sequence of Arabidopsis thaliana CCD1 (Accession: NM_116217) continued

AGATGACAGGTGGGATGTTTGGAAGGAAGGGGCAAAAGATAAAGGGGACAGTGGT GTTGATGCCAAAGAATGTGTTGGACTTCAACGCCATAACCTCCGTCGGAAAAGGCA GTGCTAAGGACACCGCCACCGATTTCTTGGGCAAAGGCTTGGACGCATTAGGTCATG CAGTTGATGCTCTCACTGCCTTCGCTGGCCATAGCATCTCCTTGCAGCTTATCAGTGC TACTCAGACTGATGGTAGTGGAAAAGGAAAAGTTGGAAACGAAGCCTATTTGGAAA AACATCTTCCGACCTTGCCAACGTTGGGAGCAAGGCAGGAAGCATTCGATATTAACT TTGAATGGGATGCTAGTTTTGGAATTCCAGGAGCATTTTACATCAAAAACTTTATGA CTGATGAGTTTTTCCTCGTCAGTGTTAAACTCGAGGACATTCCAAACCATGGAACCA TTAACTTCGTTTGTAACTCATGGGTTTATAACTTCAAAAGTTACAAAAAGAATCGCA TTTTCTTTGTCAATGATACATATCTTCCGAGTGCTACACCAGGTCCACTAGTTAAGTA CAGACAAGAAGAATTGGAGGTTTTAAGAGGAGATGGAACAGGGAAGCGCAGAGAC TTTGACAGAATCTATGATTATGATATCTATAATGATTTGGGCAATCCAGATGGTGGT GATCCTCGCCCAATCATTGGAGGCTCTAGCAACTATCCTTACCCTCGCAGGGTTAGA ACCGGTAGAGAAAAGACCAGGAAAGATCCCAACAGTGAGAAACCAGGCGAGATAT ATGTTCCAAGAGATGAAAACTTCGGTCACTTGAAGTCATCTGATTTCCTTACATATG GAATCAAATCCTTATCTCAGAACGTGATACCTTTGTTCAAATCTATAATATTGAACTT AAGGGTCACATCGAGTGAGTTCGATAGCTTCGACGAAGTGCGTGGTCTCTTTGAAGG TGGAATCAAGCTGCCAACAAATATACTGAGCCAAATTAGCCCCTTACCAGTCCTCAA Figure A-3. Sequence of Glycine max Lipoxygenase 1 (Accession: U26457)

142

(Figure A-3. continued) GGAAATCTTCCGCACTGATGGTGAAAATACCCTTCAATTTCCACCACCTCATGTAAT CAGAGTTAGTAAATCTGGATGGATGACTGATGATGAGTTTGCAAGAGAGATGATTG CTGGTGTAAATCCAAATGTAATTCGTCGTCTTCAAGAGTTCCCACCAAAAAGCACTC TTGATCCCGCAACCTATGGTGATCAAACTAGTACCATAACAAAACAACAGTTGGAG ATTAACTTGGGTGGGGTCACAGTAGAAGAGGCAATTAGTGCTCACAGATTATTCATA TTAGATTACCATGATGCATTCTTCCCGTATTTGACGAAGATAAACAGCCTACCTATT GCAAAAGCTTATGCCACAAGGACAATCCTGTTCTTGAAAGACGATGGATCTTTAAAG CCACTTGCTATCGAATTAAGCAAGCCTGCAACAGTGAGTAAAGTGGTGTTGCCTGCA ACAGAAGGTGTTGAGAGTACAATTTGGTTGTTGGCCAAGGCTCATGTCATTGTGAAT GACTCTGGTTATCATCAGCTCATAAGCCATTGGTTAAATACTCATGCAGTGATGGAG CCATTTGCCATAGCAACAAACAGGCATCTCAGTGTGCTTCACCCCATTTATAAACTT CTTTATCCTCACTACAAGGACACAATAAATATCAATGGCCTTGCTAGGCAGTCCCTG ATTAACGCAGGTGGCATTATTGAGCAAACATTTTTGCCTGGAAAGTACTCCATTGAA ATGTCATCAGTTGTTTACAAGAATTGGGTTTTCACTGACCAAGCATTACCAGCTGAT CTTGTCAAGAGAGGATTGGCAGTTGAGGATCCCTCTGCCCCACATGGTCTTCGCCTT GTGATAGAGGACTACCCTTATGCTGTTGATGGACTTGAAATATGGGATGCTATTAAG ACATGGGTCCATGAGTATGTCTCTGTGTATTACCCAACAAATGCAGCAATTCAACAA GACACTGAACTTCAAGCATGGTGGAAGGAAGTTGTGGAGAAGGGTCATGGTGACTT AAAAGATAAGCCTTGGTGGCCTAAACTGCAGACTGTGGAGGATCTCATTCAATCCTG CTCTATTATCATATGGACAGCTTCGGCTCTCCATGCAGCTGTTAATTTTGGGCAATAC CCTTATGGAGGTTATATCGTGAACCGTCCAACTCTAGCCAGAAGGTTTATCCCAGAA GAAGGAACCAAAGAATATGATGAGATGGTGAAGGATCCTCAAAAGGCATATCTGAG AACAATCACACCCAAGTTCGAGACCCTTATTGACATTTCAGTGATAGAGATATTGTC AAGGCATGCTTCTGATGAGGTCTACCTTGGCCAAAGGGATAATCCAAATTGGACTAC GGATTCAAAGGCATTGGAAGCTTTCAAAAAGTTTGGAAACAAACTGGCAGAAATTG AGGGAAAAATCACACAGAGGAACAATGATCCAAGTCTGAAAAGCCGACATGGGCC AGTTCAGCTTCCATACACATTGCTCCATCGTTCAAGTGAGGAAGGGATGAGTTTCAA AGGAATTCCCAACAGTATCTCCATCTAAAATGTGTGTGTGGTTTGCTTATCTATTGTG CTTTTGAATAAAATAGACAATACTTGTCTATGGTTATTATTGGCTGTATGTCTGTATT TGGATGCTCTCGATCGGTTTGCAAGTAATAAGAGTGTTTTCACTGTCACTTTGTATTT CGATCATCTTAATTATGTTTACTAGTAATAATGTGGAAGCTGTACGTTTGTTAATTCT AGGTTA Figure A-3. Sequence of Glycine max Lipoxygenase 1 (Accession: U26457) continued

CATATGCATTATAGTAGTGGTAATTATGAAGCTTTTGTAAACGCAAGTAAACCTAAG GATGTCGATCAGAAGTCCGCATATCTTGTTGGTTCAGGTTTGGCATCGCTTGCTAGT GCTGTATTTTTAATTCGTGATGGTCACATGAAGGGTGATAGAATTCATATCCTTGAA GAATTGAGCCTTCCAGGTGGTTCAATGGATGGGATCTATAATAAGCAAAAAGAAAG CTACATCATTCGTGGTGGTCGTGAAATGGAAGCCCATTTTGAATGCTTGTGGGACTT GTTTAGATCGATTCCATCAGCTGAAAATAAAGATGAATCGGTCCTGGATGAATTTTA CCGTTTAAATAGAAAAGATCCAAGTTTCGCAAAGACTCGTGTCATTGTTAACCGCGG ACATGAACTTCCAACTGACGGTCAATTACTTCTTACTCCCAAGGCTGTTAAAGAAAT TATTGATCTTTGCTTAACTCCTGAAAAAGATTTACAAAATAAAAAAATTAATGAAGT Figure A-4. Sequence of Lactobacillus acidophilus strain KCTC Lineolate 13-Hydratase (Accession: KJ560553.1)

143

(Figure A-4. continued) CTTTAGTAAAGAATTTTTTGAATCAAACTTCTGGCTTTACTGGTCAACGATGTTTGCC TTTGAGCCATGGGCAAGTGCGATGGAAATGCGTCGTTACTTAATGCGTTTTGTTCAA CACGTTTCTACACTTAAGAATTTATCATCACTACGCTTTACTAAGTATAACCAATATG AATCATTAATTTTACCAATGGTTAAATACTTGAAAGATCGCGGCGTGCAATTCCATT ACAACACCGTTGTTGATAATATCTTTGTTAACCGTTCAAATGGTGAAAAGATTGCTA AGCAAATTCTTTTAACTGAAAACGGTGAAAAAAAGAGCATCGATTTAACAGAAAAT GACCTCGTCTTCGTTACTAACGGTTCAATTACTGAAAGTACAACTTATGGTGATAAC TTGCACCCAGCTTCTGAGGAACATAAATTAGGTGCTACTTGGAAATTATGGCAAAAC TTGGCAGCGCAAGATGATGACTTCGGTCACCCAGATGTCTTCTGCAAGGATATTCCA AAGGCTAACTGGGTAATGTCTGCTACAATTACTTTTAAGAATAATGATATTGTGCCA TTCATTGAAGCAGTTAATAAGAAGGACCCACACAGCGGCTCAATTGTAACTAGTGG GCCTACTACGATTAAGGATTCTAACTGGCTACTTGGTTATTCAATCAGTCGTCAGCCT CACTTTGAAGCACAAAAGCCTAACGAATTGATTGTATGGCTTTATGGTTTGTTCTCA GACACCAAAGGTAACTATGTTGAAAAGACTATGCCTGACTGTAACGGTATTGAATTA TGTGAAGAATGGCTTTACCACATGGGTGTTCCTGAAGAAAGAATCCCAGAAATGGC TTCAGCTGCTACGACTATTCCAGCACACATGCCATATATTACTTCATACTTCATGCCA AGAGCATTAGGCGACAGACCCAAGGTTGTGCCAGACCACTCAAAGAACTTGGCCTT CATTGGTAACTTTGCTGAAACGCCAAGAGACACTGTCTTTACCACTGAATACTCTGT CAGAACTGCGATGGAAGCTGTATACACCTTGCTTAACATTGATCGTGGTGTGCCAGA AGTATTTGCATCTGCCTTCGATGTCAGAATGCTCATGAACGCAATGTACTACTTGAA TGATCAAAAGAAGCTTGAAGATCTTGATTTGCCTATTGCTGAAAAGTTGGCAATTAA GGGGATGCTCAAGAAAGTTAAGGGCACTTATATAGAGGAATTGCTTAAGAAGTATA AGTTGGTTTAGGGATCC Figure A-4. Sequence of Lactobacillus acidophilus strain KCTC Lineolate 13-Hydratase (Accession: KJ560553.1) continued

Plasmid Maps

Figure A-5. Plasmid map of pEA1

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Figure A-6. Plasmid map of pEA2

Figure A-7. Plasmid map of pEA3

Figure A-8. Plasmid map of pEA4

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Figure A-9. Plasmid map of pEA6

Primer Sequences

Table A-1. List of Sequence Altering Primers for AtCCD1

Mutation Sequence NdeI→ EcoRI Fwd (pEA1) 5’-CACTGAGAATTCGCTGCTAACAAAGCCCG-3’ NdeI→ EcoRI Rev (pEA1) 5’-AGCAGCGAATTCTCAGTGGTGGTGGTGGTG-3’

Table A-2. List of Mutagenic Primers for AtCCD1

Mutation Sequence AtCCD1 F98A Fwd 5’-ATATCACTGGGCGGATGGAGATGGGATGATT-3’ AtCCD1 F98A Rev 5’-CATCTCCATCCGCCCAGTGATATCCAGCG-3’ AtCCD1 M269A Fwd 5’-AGAGCCTATCGCGATGCATGATTTTGCTATTACTGAG-3’ AtCCD1 M269A Rev 5’AAATCATGCACGCGATAGGCTCTGATATAGTAATTGGG-3’

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APPENDIX B FATTY ACID STRUCTURES

Table B-1. Significant chemical structures

Substrate Structure

β-Apo-8’-Carotenal

Linoleic Acid

Methyl Lineolate

Linolenic Acid

Arachidonic Acid

Eladic Acid

Palmitoleic Acid

Oleic Acid

Erucic Acid

Lauric Acid

Hexanoic Acid

Valeric Acid n-Hexanal n-Heptanal

E,E-2,4-Decadienal

E-4-Decenal 1-Octene

Trans-Crotonic Acid

147

Table B-1. Continued

9E, 11E CLA

9Z, 11E CLA

10E, 12Z CLA

Linoeladic Acid

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APPENDIX C ADDITIONAL GC/MS DATA

GC Chromatograms

Figure C-1. GC chromatogram for lipoxygenase biotransformation products

Figure C-2. GC chromatogram for lineolate 13-hydratase biotransformation products

149

Mass Spectra

Figure C-3. Mass spectrum analysis of linolenic acid hydration with lineolate 13-hydratase

Figure C-4. Mass spectrum analysis of palmitoleic acid hydration with lineolate 13-hydratase

150

Figure C-5. Mass spectrum analysis of methyl lineolate hydration with lineolate 13-hydratase

Upon speaking with the mass spec faculty, the methyl group on the carboxylic acid will ionize off resulting in an MSTFA derivitized m/z 371 as a major peak.

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APPENDIX D NMR DATA

Figure D-1. 1H NMR of DNPH derivitized n-hexanal

Figure D-2. 1H NMR of pure n-hexanal

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BIOGRAPHICAL SKETCH

Erica Amato Simmons was born in Indianapolis, IN in 1991. She graduated from

Northern Kentucky University in 2013 with a Bachelor of Science in chemistry. Upon graduation she moved to the University of Florida to join the biochemistry Ph.D. program. There she joined the laboratory of Dr. Jon Stewart and was given the project of converting linoleic acid into n- hexanal through an all-natural process. Her research interests include merging organic chemistry and biocatalysis in medical chemistry applications. She expects to earn her Ph.D. by December of 2017.

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