BIOCHEMICAL CHARACTERIZATION OF N-TERMINALLY TAGGED STYRENE MONOOXYGENASE FROM PSEUDOMONAS
A thesis submitted to the faculty of San Francisco State University AS In partial fulfillment of The Requirements for The Degree
Master of Science In • O U Chemistry: Biochemistry
by
Nonye Nwa-Niaf Okonkwo
San Francisco, California
May 2015 Copyright by Nonye N. Okonkwo 2015 CERTIFICATION OF APPROVAL
I certify that I have read The Biochemical Characterization of N-Terminally Histidine
Tagged Styrene Monooxygenase from Pseudomonas by Nonye Nwa-Niaf Okonkwo, and that in my opinion this work meets the criteria for approving a thesis submitted in partial fulfillment of the requirements for the degree: Master of Science in Chemistry: Biochemistry at San Francisco State University.
Professor of Biochemistry
Weiming Wu, PhD Professor of Biochemistry « BIOCHEMICAL CHARACTERIZATION OF N-TERMINALLY TAGGED STYRENE MONOOXYGENASE FROM PSEUDOMONAS
Nonye Nwa-Niaf Okonkwo San Francisco State, California 2015
Flavoprotein monooxygenases are involved in a wide variety of biological oxidations due to their ability to utilize organic flavin cofactors as substrates to stereo- and regioslectively produce valuable bioactive compounds. Metabolism of styrene by Pseudomonas putida (S12) bacteria is accomplished by a two-component flavoenzyme system, which consists of styrene monooxygenase A (SMOA) and styrene monooxygenase B (SMOB), a reductase. Our present research evaluates the catalytic mechanisms of an N-terminally histidine-tagged version of the styrene monooxygenase reductase (NSMOB) component. Fluorescence monitored titrations at 4°C confirmed the equilibrium dissociation constant of NSMOB to have a Kd value of ~ 50 nM, an order of magnitude greater than the wild-type reductase. Steady-state kinetic analysis at 30°C also determined that a double-displacement mechanism with NADH as the leading substrate is the preferred method of FAD reduction. Due to the significant change in FAD binding affinity and catalytic mechanism, stopped-flow fluorescence and absorbance was used to evaluate the pre-steady state kinetics of NSMOB. We determined that the hydride-transfer rate constant is k = 48 s'1, which is identical to the wild-type enzyme at k = 50 s'1 at 15°C, which effectively resolves the rate-limiting step for the N-terminally tagged enzyme. These findings will be presented together with their implications for the engineering of N-terminally tagged enzymes and N-terminally linked fusion proteins as biocatalysts for the production of essential chiral epoxides.
I certifWhat the Abstract is a correct representation of the content of this thesis.
Date PREFACE AND AKNOWLEDGMENTS
I would like to thank Dr. George Gassner for four years of exceptional mentorship and outstanding research experience. I would like to recognize Berhanegerbial Assefa for his preliminary work on the N-terminally tagged SMOB. Additionally, I would like to recognize the San Francisco State Women’s Association, College of Science and Engineering, PG&E,
CSU Sally Casanova Pre-Doctoral Program and Undergraduate NIH-MARC and Graduate
NIH-RISE MBRS fellowships for their continued financial support and professional development assistance. This work was supported by NIFISC1 GM081140 to George
Gassner and Nonye Okonkwo was supported by the NIH-MARC 5T34-GM008574 and NIH-
MBRS RISE fellowships.
v TABLE OF CONTENTS
List of Tables...... viii
List of Figures...... ix
List of Equations...... xi
List of Appendices...... xii
Introduction...... 1
1.1 Significance and toxicology of styrene in the environment...... 1 1.2 Microbiology of styrene degradation...... 5 1.3 Styrene catabolic and detoxification pathway...... 11 1.4 One- and two-component styrene monooxygenases...... 14 1.5 Catalytic mechanisms of styrene monooxygenases...... 18 1.6 Styrene monooxygenase structure...... 25 1.7 Naturally-occurring and artificially engineered fusion proteins...... 27 1.8 Putative complexes...... 30 1.9 Transition to thesis research...... 32
Methods...... 34
2.1 Protein expression and purification...... 34 2.2 Activity and purity assays...... 36 2.3 Time-dependent activity studies...... 39 2.4 Steady-state kinetic analysis ofNSMOB...... 40 2.5 Estimation of equilibrium dissociation constants...... 43 2.6 Pre-steady-state kinetics...... 44
Results...... 46
3.1 Flavin-refolded SMOB experiments with FAD, FMN, and riboflavin....46 3.2 Flavin-refolded SMOB time dependence studies...... 52 3.3 NSMOB mechanistic studies...... 56 3.4 Estimation of equilibrium dissociation constants by fluorescence Monitored Titrations...... 60
vi 3.5 NSMOB single-turnover studies...... 61 3.6 Efficiency of oxidative- and reductive half-reactions...... 65
Discussion...... 68
4.1 N-terminal tags and implications in protein engineering...... 68 4.2 Isoalloxazine ring environment: SMOB refolding and stability...... 69 4.3 SMO Rate-limiting hydride-transfer step ...... 72 4.4 SMO Flavin-transfer mechanisms and protein-protein interactions...... 75 4.5 Engineered Flavin Monooxygenases as Efficient Biocatalysts...... 77
References...... 78
Appendix...... 84 LIST OF TABLES
Table Page
1. Flavin-refolded SMOB total protein and specific activity...... 50
2. Comparison of NSMOB Kinetic Parameters...... 57
3. Hull Hypothesis Comparison of BiBi Sequential and Double-Displacement Mechanisms...... 59 LIST OF FIGURES
Figure Page
1. Chemical Structure of Styrene and (S)-Styrene-7, 8- oxide...... 4
2. Metabolic fate of styrene...... 7
3. Bacterial degradation pathway for styrene...... 9
4. Styrene monooxygenase catalysis...... 12
5. C4a-hydroperoxyflavin intermediate...... 14
6. Oxidative and reductive half-reactions...... 18
7. Isoalloxazine catalytic cycle of flavin-dependent monooxygenases...... 23
8. N-terminally tagged SMOB...... 25
9. Overall structure of N-terminally tagged styrene monooxygenase A ...... 26
10. Organization and mechanism of SMO systems...... 28
11. Possible complex formation of native and N-terminally tagged SMOB and SMOA during the FAD-exchange reaction...... 32
12. Flavin-refolding process of SMOB in 8M urea...... 46
13. Flavin structure: riboflavin, FMN and FAD...... 47
14. Flavin-refolded SMOB SDS-PAGE...... 49
15. Graph of flavin-refolded SMOB total protein assay results...... 50
16. Flavin-Refolded Kinetic Activity Assay...... 51 17. Flavin-re folded SMOB Kinetic Activity Graphs...... 51
18. Half-life of SMOB in the presence of FAD, FMN, and riboflavin...... 52
19. Absorbance flavin reconstituted With FAD at -20°C and 4°C...... 53
20. Fluorescence spectra of flavin reconstituted with FAD...... 54
21. Temperature-dependent Estimation of fluorescence dissociation constants by fluorescence monitored titrations...... 55
22. Ribbon structure of N-terminally tagged FAD-bound SMOB...... 56
23. Global fit NSMOB double-displacement mechanisms...... 58
24. Fluorescence titration monitoring of FAD to NSMOB...... 60
25. Steady-state NSMOB kinetic data...... 62
26. Time averaged single-turnover NSMOB stopped-flow data...... 63
27. Absorbance and fluorescence measurements of hydride-transfer reaction...... 64
28. Styrene epoxidation in the reaction catalyzed by NSMOB and NSMOA...... 65
29. Kinetics of FAD reduction in the reaction of SMOB in the presence ofNADH, FAD, SMOA and oxygen...... 66
30. Interchange of Sequential and Double Displacement Reaction Mechanisms based on Kd...... 70
31. Native SMOB catalytic mechanism...... 73
32. N-terminally tagged SMOB catalytic mechanism...... 73
x LIST OF EQUATIONS
Equation Page
1. Velocity rate equation for time dependence studies...... 40
2. Kinetic fitting equations: V, Vmax and Km apparent...... 41
3. Ordered sequential mechanism equation...... 42
4. Double-displacement mechanism equation...... 43
5. FAD-binding constant quadratic equation...... 44
6. Steady-state fitting equation: exponential + linear...... 62 LIST OF APPENDICES
Appendix Page
1. FPLC fraction plot...... 84
2. N-SMOB SDS-PAGE analysis...... 85
3. NSMOB BSA standard curve...... 86
4. NSMOB total protein...... 87
5. Steady-state kinetic plots ofNSMOB with NADH and FAD...... 88
6. Non-time averaged single-turnover NSMOB stopped-flow data...... 89 1
INTRODUCTION
1.1 Significance and toxicology of styrene in the environment
Our modern society is extremely dependent on the mass-production of resources involving the release of artificially synthesized polymers and natural elements that may be harmful to microorganisms and entire ecosystems (Donner, 2010). Because of our increasing reliance on the many important industrial processes and facilities that produce these polymers and compounds, our environment is continuously polluted with the byproducts of many known and unknown xenobiotic materials. Quantification of the level and expanse of xenobiotics released from different sources, both industrial and natural, into the environment has been challenging, but it is imperative to the management of toxic wastes and understanding the difference between beneficial xenobiotic sources and limiting our exposure to putative carcinogens (Donner, 2010).
During the last century extensive population growth and rapid industrialization has caused an increase in the number and type of xenobiotic compounds present in our natural environment (Donner, 2010). The term xenobiotic refers to any substance that is foreign to biological systems, but can be found in them although not produced by them (Donner,
2010). Exogenous xenobiotics can gain entry into host organisms through the diet, pharmacological drugs, or direct transfer, whereas endogenous products are synthesized in the body as metabolites of various biological processes (Arora, 2009). Artificially synthesized compounds and naturally-occurring elements can be placed in this category, 2
and are now present in unnaturally high concentrations due to increased use of anthropogenic processes, which in turn amplifies their likelihood of altering the physical, chemical, and biological properties of the environment over time (Donner, 2010).
Xenobiotic compounds are generally metabolized by the conversion of the lipophillic compound into a hydrophilic product that can be easily excreted by the organism. The chemical properties and quantity of the xenobiotic compound determines its relative toxicity and persistence in the environment (Donner, 2010). Cytochrome p450 oxygenases are a large class of hemoproteins that act in a variety of enzymatic processes, most importantly drug and toxin metabolism, therefore acting as the chief gatekeeper in the biotransformation of harmful xenobiotic compounds (Donner, 2010). This and other related degradation pathways are of tremendous significance in environmental science because humans and microorganisms may or may not possess the ability to break down a pollutant; the increased public awareness about the hazards and toxicity of these compounds has encouraged the development of new versatile technologies for bioremediation (Arora, 2009).
Millions of dollars are being spent to fully understand the breadth of potential bioremediation applications due to commercial mass-production and the health impact of xenobiotics due to their abundant negative physio-chemical activities and positive roles in drug discovery (Arora, 2009). For example, 95% of artificially synthesized 1, 2- dichloroethane is used to produce vinyl chloride, which is then used to make polyvinyl chloride (PVC) (Arora, 2009). PVC is presently the third most produced synthetic polymer 3
after polyethylene and polypropylene, and during the 70’s it was determined that vinyl chlorides were linked to increased incidence of cancer in tire plant workers, which led to its decreased use in healthcare and occupations requiring long-term human exposure
(Arora, 2009).
In light of the increased regulation ofxenobiotic compounds produced industrially, research has been geared towards fully understanding the genotoxicity of many widely used polymers. Styrene is an essential plastics monomer used worldwide; it is used to make rubber, plastics, and polystyrene copolymers by commercial industries at the rate of fifteen billion pounds every year in the United States alone (IARC, 1994). The general public is directly exposed to styrene through contaminated water and inhalation (-2% of pulmonary uptake), with skin absorption being very low on the exposure charts (Henderson, 2005).
Furthermore, 89 % of inhaled styrene is absorbed by the blood where it has a Vi life of 41 min and 32-46 hours in fatty tissues (Guillemin and Berode 1988). Airborne styrene interacts with the trophosphere layer of the atmosphere with a V2 life of about 2.5 hours; biodegradation of styrene in soil occurs at about 87 - 95% after about 16 weeks (US
Inventory of Toxic Compounds 2001).
The majority of recent ecological efforts are now aimed at targeting the quantification and degradation of polymers in aquatic and land environments through population studies, which will help understand possible the genotoxic events in xenobiotic mixtures (Arora, 2009; Vodicka, 2002). Styrene is and its immediate metabolite, styrene-
7, 8- oxide (SO) have been classified as human carcinogens based on extensive research in 4
animals (Figure 1) (IARC, 1994). Some of the first studies onto 1,3-butadiene and styrene occupational exposure among workers in the synthetic rubber industry correlated the
incidence of leukemia mortality (Macaluso, 1996), and increased excretion of mandelic
acid (MA) and phenylglyoxylic acid (PGA) in the urine of exposed subjects (IARC, 1994).
The most disturbing genotoxicity results implicated styrene in DNA strand breaks, DNA
alkylation, and tumorgenesis in rats (Lutz, 1993). Additionally, styrene oxide-DNA and
styrene oxide-albumin adducts were also found in high concentration in the blood of
plastics workers (IARC, 1994), underscoring the importance of elucidating the biochemical
degradation pathways for harmful xenobiotics including styrene and styrene oxide.
Due to the associated health risks and ubiquitous nature of styrene in industrial
production, it also represents an environmental contaminant as it is present in food items, tobacco smoke, and engine fumes (IARC, 1994). o Biologically styrene acts as a human cellular V
membrane disrupter and manifests its intracellular I effects through cytochrome p450 isofroms, which
convert styrene to SO and 4-vinylphenol, which Styrene Styrene-7, 8- Oxide Figure 1: Chemical Structure of have biological activity as carcinogens and Styrene and (Sj-Styrene-7, 8- Oxide. pulmonary and hepatic toxins (IARC, 1994).
Current methods for alleviating the mass pollution of styrene include land spreading,
underground injection, and combustion to create usable energy (Westblad, 2002).
Improving our understanding of the mechanisms by which microorganisms detoxify their 5
habitats is essential in providing the basis for elucidating the strategies used by the natural world to cope with the increasingly contaminated environment.
1.2 Microbiology of styrene degradation
Styrene is named after Storax balsam, a resin produced by the Liquidambar trees, a product that is naturally found in some plants and foods including cinnamon, coffee beans, and peanuts (Vodicka, 2002). Styrene is a simple alkylbenzene, an unsaturated aromatic monomer, and is also known as phenylethylene, vinylbenzene, styrol, and cinnamene (Mooney, 2006). Laboratory classifications identify styrene as a sweet smelling, colorless, oily liquid known to undergo spontaneous polymerization at room temperature (Mooney, 2006). Large scale production is achieved by the direct dehydrogenation of ethyl benzene, a process that comprises 85 % of its commercial synthesis (Mooney, 2006).
The first step of styrene metabolism forms the highly bioreactive styrene -7, 8- oxide via hepatic microsomal cytochrome p 450 monooxygenases in humans with a small fraction of this reaction occurs in the lungs (Hartmans, 1990). SO plays a vital role in styrene toxicity, and serves as the primary source of styrene’s carcinogenic effects because its electrophillic nature allows it to effectively covalently bind biological macromolecules
(Wu, 2011). Next, SO is hydrolyzed to styrene glycol by microsomal epoxide hydrolases and then oxidized via alcohol dehydrogenases to form its urinary metabolites, mandelic acid (MA) and phenylglyoxylic acid (PGA). MA and PGA represent 95% of styrene urinary excretions in humans, and subsequent transamination reactions can convert these 6
metabolites to phenylglycine amino acid (Reuff, 2009). In humans, urinary analysis of the primary MA and PGA metabolites are used as biomarkers to measure styrene exposure through work or pollution (Reuff, 2009). Along with this knowledge much of the current research on the microbiology of styrene degradation has been directed at understanding human susceptibility to genetic damage at the hands of SO mediated by cytochrome p450 enzymes (Henderson, 2005).
In vivo styrene is metabolized by cytochrome p450 monooxygenases, including the major isoforms: CYP1A1 and CYP2E1. The CYP2E1 class of p450 enzymes are responsible for the biotransformation of styrene to SO, and polymorphic differences have been shown to influence breakdown and mediate SOs genotoxicity, because the first step in the microbiology of styrene degradation is the functional binding of styrene to CYP2E1
(Figure 2) (Wu, 2011). Human CYP2E1 possesses a substrate binding region with six amino acids: phenylalanine-298, alanine-299, glycine-300, threonine-301, glutamic acid-
302 and threonine-303 and cysteine-437 interfaces with the heme group, all residues are conserved in mammals and bacteria (Wu, 2011). Styrene dimers, trimers, and pentamers were investigated for their CYP2E1 affinity and binding energy experiments indicated that styrene dimers may dock more efficiently to cytochrome p450 (Wu, 2011). Additionally,
CYP2F2 was also found to be responsible for SO production and the expression levels of both isomers are correlated to its physiological and genotoxic effects (Wu, 2011). 7
HsCj,
Myrono 3,4-o*kJ«
Glticuronicta imd sutfato co n ju g a te *
i\yiom /.8-ft*iCfc <80)
Figure 2: Metabolic Fate of Styrene. Cytochrome P450 enzymes catalyze the oxidation of styrene styrene-7, 8-oxide (SO). SO can be hydrolyzed to styrene glycol by the microsomal epoxide hydrolase, and subsequently oxidized by alcohol and aldehyde dehydrogenases to the main urinary metabolites, mandelic acid (MA) and phenylglyoxylic acid (PGA) (major pathway). The minor metabolic pathway involves the conjugation of SO with glutathione (GSH) via glutathione S-transferase (GST). (Reuff, 2009)
The remaining 1% of styrene can participate in two minor degradation pathways involving: glutathione (GSH) and glutathione S-transferase (GST) or a 3, 4 ring epoxidation reaction. This small fraction is converted to SO and then conjugated with GSH via GST, and the residual metabolites are converted to 4-vinyl phenol, which is then excreted in the urine (Reuff, 2009). GST is a styrene inducible enzyme involved in the metabolism of a wide variety of xenobiotic epoxide compounds and retains its substrate specificity dependent on its cytoplasmic classification (Reuff, 2009). The most prevalent iso form, GST-P1 is found in the lungs and has been linked to styrene’s downstream carcinogenic affects upon inhalation of tobacco; relating its importance to the clearance of toxicants and pro-carcinogens, moreover, overexpression of GST enzymes has been linked to tumors and drug resistant cell lines (Reuff, 2009).
1.2.1 Regulation of microbial styrene degradation
Styrene and its intermediate SO are known xenobiotic carcinogens and the enzymatic epoxidation of aromatic compounds represents a versatile supply of readily available and cheap substrates in pharmaceutical synthesis (Reuff, 2009). The biotransformation of styrene to SO bacterial monooxygenase enzymes are of particular interest due to their versatility and exceptional enantioselectivity (Montersino, 2011;
Huijbers, 2014). Microbial communities have survived long-term environmental exposure to styrene and boast the necessary catalytic mechanisms needed to detoxify harmful aromatic compounds that most mammals do not have.
Research to date on the regulation of microbial styrene degradation is concentrated on the genetic characterization of the catabolic operons together with functional analysis of key enzymes from the pathways utilizing one- and two component monooxygenases
(O'leary, 2002). This new direction has provided valuable information on the organization of styrene metabolic genes and future research will focus on understanding the physiological factors and unique environmental conditions that influence the expression of the Pseudomonas sp. vital gene products (Otto, 2004). 9
styC styD styrene styrene oxide phenylacetaidehyde monooxygenase dehydrogenase styrene styrene oxide phenylacetaidehyde
OH ^ ^ ^SCoA TCA Cycle O phenylacety! coA ttgase phenylacetic acid phenylacetyl coenzyme A
Figure 3: Bacterial degradation pathway for styrene, showing intermediates, enzyme names and corresponding genes (Mooney, 2006).
Previous studies on microbial catabolic mechanisms were concentrated on the characterization of genetic operons from various strains (Beltrametti, 1997). It was demonstrated that styrene degradation by Pseudomonas sp. bacteria proceeds via side chain oxidation utilizing an upper and lower pathway (O’leary, 2001). The upper pathway involves styrene, styrene oxide, and phenylacetic acid (PAA); the lower pathway begins with PAA (O’leary, 2001). Subsequent genetic studies have identified the genes involved in the upper pathway conversion of styrene to PAA in number bacteria that are able to detoxify styrene, most notably the Pseudomonas fluorescens, Pseudomonas putida, and
Rhodococcus opacus species.
The upper pathway is the most commonly described route for styrene degradation and involves the complete oxidation of styrene to form PAA, which is then converted to
TCA cycle intermediates illustrated in Figure 3 (Mooney, 2006). The epoxidation of the 10
styrene vinyl side chain is catalyzed by a two-component styrene monooxygenase, encoded by the styA and styB genes. StyA possesss styrene monooxygenase activity and converts styrene to SO utilizing electrons from FADH2 ; styB retains FAD reductase activity and transfers electrons from NADH to FAD+ to supply FADH2 for sty A. (S)- styrene oxide is then isomerized by the styC gene product, styrene oxide isomerase (SOI), to yield PAA.
Also present on the operon is styD, which encodes the phenylacetaldehyde dehydrogenase
(PADH) and necessary for the oxidation of phenylacetaldehyde to phenylacetic acid
(Mooney, 2006). In the lower pathway phenylacetic acid is ligated to coenzyme A to produce phenylacetyl coA, via phenylacetyl coA liase, which is encoded by paaF2, and then hydrolyzed to produce acetyl coA, which enters the tricabrocylic acid cycle (TCA)
(Teufel, 2010).
Other important genes, styS and styR, are associated with the modulation of styrene catabolism genes and sequence homology analyses suggest that they are involved in other two-component regulatory systems found in both prokaryotic and eukaryotic species
(Mooney, 2006). The styS gene product shows similarities to sensor kinase proteins that generally regulate aerobic and anaerobic metabolism of toluene in P. putida FI bacteria
(Mooney, 2006). StyR belongs to the FixJ/NarL subfamily and acts as a response controller with homology to the regulators of most two-component systems (Mooney, 2006). Studies using gel retardation and DNase I footprinting experiments found that styS and styR are essential for controlling the upper pathway, involving styrene, SO and PAA (O’leary, 2001). 11
The activation of the main upper pathway is dependent on styrene, and has the ability to be shut-off in the in the presence of other carbon sources like phenylacetic acid or citrate (O’leary, 2001). Pseudomonas putida CA-3 displays styrene detoxification pathway repression in the presence of citrate through a process that involves the reduced transcription of styS, styR and styA genes as long as the catabolite is present (Otto, 2004).
Also, the restriction of inorganic nutrients like: phosphate, sulfur, and nitrogen have similar repressive effects on P. putida bacteria making them very sensitive to catabolite repression.
It is clear that a variety of factors act in concert to respond to environmental stimuli during styrene degradation however, the importance and roles of all styABCD gene products are still in need of more stringent characterization (O’leary, 2002). Understanding the complex flavin-dependent mechanisms that styrene monooxygenase employs to biologically oxidize organic compounds has a broad impact on the various potential applications of this biocatalyst in medicine and technology (Sucharitakul, 2014).
1.3 Styrene catabolic and detoxification pathway
The new millennium witnessed a significant increase in the global production and utilization of alkylbenzene derivatives as polymer-processing industries became major contributors to the pollution of natural resources through the discharge of styrene- contaminated effluents and off-gases (Otto, 2004). Toxicologists are concerned with human exposure to styrene and scientific research is interested in understanding the regulation of early catabolic intermediates, which are known to have a wide variety of carcinogenic health affects (Vodicka, 2002). These aspects have prompted investigators to 12
identify routes of styrene degradation in microorganisms, given the potential application of these organisms in bioremediation strategies and pharmacology (Mooney, 2006).
Optically active epoxide compounds are of industrial and medical importance for their use as precursors for the chemical synthesis of drugs thereby putting the biocatalysts that are able to synthesize pure styrene oxide at the center of attention (O’leary, 2001). The styA and styB encoded styrene monooxygenase (SMO) enzyme is a well characterized two- component flavoprotein that catalyzes the conversion of styrene to styrene-7, 8- oxide.
Along with its impeccable regio- and enantioselectivity it can efficiently produce both the
(S)- and (R)- SO enantiomers depending on the microbial strain (Panke, 2000).
Two-component styrene
monooxygenases, members of the class E
flavoprotein monooxygenases, are able to
catalyze the stereospecific epoxidation of
vinyl benzene derivatives to create vital
bioactive compounds (Van Berkel, 2006;
Montersino, 2011). The first step of the
Figure 4: Styrene Monooxygenase styrene detoxification pathway in (SMO) Catalysis. SMOB (pink) catalyzes NADH-specific reduction of Pseudomonas sp. converts styrene to (S)- FAD and SMOA (green) performs subsequent FAD-dependent styrene-oxide by using an NADH-dependent epoxidation of styrene to form (S)- styrene oxide (Hartmans, 1990). reductase (SMOB) and an FAD-specific monooxygenase (SMOA) (Van Berkel, 2006; Montersino, 2011) as seen in Figure 4. The 13
oxidation efficiency of styrene monooxygenase involves the transfer of reduced FAD from the SMOB reductase component to the active site of SMO A in the presence of molecular
oxygen (O2) and the stabilization of the FAD C4a-hydroperoxyflavin (Kantz, 2005; Kantz,
2011; Morrison, 2013). The reactive oxygen species then stereoselectively attacks the
styrene vinyl group to catalyze the epoxidation and formation of an FAD C4a-hydroxide,
which dehydrates to reform oxidized FAD (Kantz, 2011).
SMOs versatility is also demonstrated in its ability to catalyze the
monooxygenation of aromatic substrates including the conversion of indole to indole
oxide, and indene to indene oxide (Hollman, 2003). Indene oxide is a vital precursor of the
anti-HIV-1 drug, Crixavin- cis-lS, 2R-aminoindanol, an intermediate in the drug synthesis.
Laboratory synthesis of indene oxide is very difficult, and in addition to possessing the
machinery needed to oxygenate lipophillic compounds, P. putida are affected by a plethora
of extracellular and intracellular conditions (O’leary, 2002).
1.3.1 Factors mediating SMO catalytic efficiency
The enzyme ratio of styA to styB has been shown to significantly influence the rate
of epoxidation to yield SO (Otto, 2004). The highest yields of styrene biotransformation
are achieved when molar amounts of the SMOB reductase were equal or higher than that
of the SMOA epoxidase; this also holds true for some two-component systems however,
the impact of styA and styB expression is diverse depending on the microbial species (Otto,
2004). For the SMO system, reduced FAD is limiting in the SMOA catalyzed reaction, so 14
the addition of more SMOB reductase is expected to increase the overall intracellular concentration of FAD and enhance all its dependent reactions (Otto, 2004).
Conversely, when SMOA concentrations exceed that of the reductase, the system becomes uncoupled causing FAD to react with O2 to yield hydrogen peroxide and superoxide, harmful reactive oxygen species (Kantz, 2005), which may present a challenge in catalytic efficiency for the development of biocatalysts representing both the two- component and one-component monooxygenase families (Tischler, 2013). In light of these limitations, new research has uncovered vital details about the method of flavin reduction and how various non-active and active-site characteristics effect the speed and delivery of reduced FAD to the epoxidase component, a similar challenge for all two-component flavin
monooxygenases (Lin, 2012). - i A R' ' - * 1.4 One- and two-component
id\ /J monooxygenase systems R ' oh H !I . OH° H XX^v™ M Flavin-dependent H O if R R i / HO ^ vn J \{4) enzymes are highly abundant in / \ nature and are widely regarded 1 X R R’ ______x-s,s,se.n,ft______as indispensable tools in the Figure 5: C4a-hydro-peroxyflavin Intermediate Acts as a Precursor for Many Important Reactions synthesis of biologically active Catalyzed by Flavin-Dependent Monooxygenases. (1) Epoxidation of C=C double bonds, (2) Baeyer- compounds (Chaiyen, 2012). Villiger oxidation of ketones, (3) hydroxylation of phenols, (4) heteroatom oxygenation, (5) aldehyde The remarkable enantio oxidation, (6) halogenation reactions (Holtmann, 2014). selectivity of the one- and two- 15
component flavoprotein monooxygenases has led these enzymes to be a primary target for biotechnology due to their ability to activate molecular oxygen and the presence of an organic flavin cofactor (Chaiyen, 2012). The resulting chiral and achiral, oxygenated and hydroxylated products are high value precursors in medicinal chemistry, but the large-scale synthesis of these relevant substrates via organic chemistry is very difficult, making the application of biocatalysts in these transformations highly sought after (Bornscheuer,
2012). Furthermore, the C4a- hydroperoxyflavin acts as the active species leading to the many diverse reactions that are facilitated by flavin-dependent monooxygenases shown above in Figure 5.
Flavoprotein monooxygenases catalyze a wide range of oxygenation reactions that include hydroxylations, epoxidations, Baeyer-Villiger oxidations and sulfoxidations; the specific type of oxygenation and selectivity depends on the shape and chemical nature of the active site of each specific monooxygenase (Van Berkel, 2006; Kadow, 2014). In particular the external two-component Bayer-Villager Monooxygenases, 4- hydroxyphenylacetate 3-monooxygenase, and styrene monooxygenase are being studied extensively for their potential as marketable biocatalysts as they efficiently employ reduced
NAD(P)H as a source of electrons for their non-covalently bound FAD or FMN coenzymes
(Kim, 2008; Torres Pazmino, 2010). In general, this research is concerned with the reductases that catalyze the electron-transfer between organic flavin reductants and electron acceptors, and the oxygenase components that utilize the reductant and molecular oxygen as co-substrates. Currently, the selectivity and presence of an organic co factor has 16
garnered industrial interest in flavin enzymes as environmentally friendly oxidative biocatalysts that can perform regio- and enantioselective oxidations for the production of high-value anthropogenic chemicals and pharmaceutical intermediates (Reetz, 2013).
The classification of external flavoenzyme monooxygenases is been based on the type of chemical reaction they catalyze, redox cofactor, and sequence homology; to date six subclasses A-F have been described primarily based protein structural features (Van
Berkel, 2006). Class A and B enzymes are NAD(P)H dependent and catalyze the ortho- or para hydroxylation of aromatic compounds containing an activating hydroxyl or amino group (Van Berkel, 2006). These hydroxylases are distinct from the cytochrome p450 enzymes because they maintain the ability to hydroxy late aliphatic and aromatic compounds lacking activating functional groups (Van Berkel, 2006).
One-component monooxygenase systems are unique in that the oxidized and reduced flavin molecule resides in the same active site throughout the catalytic cycle, which protects the reduced flavin from non-specific reactions involving oxygen that cause the production of hydrogen peroxide; these non-specific reactions lead to the wasteful over consumption ofNAD(P)H energy sources (Van Berkel, 1995). Of this class, the single component, oxidoreductase 4-hydroxybenzoate 3-monooxygenase of the Pseudomonas sp. is the most well understood as an NADPH dependent protein with an N-terminal sequence indicative of a Rossmann fold, which is known to bind the ADP moiety of FAD with high affinity (Van Berkel, 1995). Their regioselective electrophillic aromatic substitution 17
mechanism includes the stabilization of a C4a-hydroperoxyflavin that interacts with nucleophillic hydroxyl or amino functional groups (Van Berkel, 2006).
Similarities among this broad class of flavoproteins are highlighted by the para hydroxybenzoate hydroxylase (PHBH) enzyme family that features a special hydrogen bond network to ensure sufficient substrate nucleophillicity for deprotonation (Van Berkel,
1995). Pseudomonasfluorescens microbes utilize a non-covalently bound FAD to catalyze the conversion o f/;—h_\ droxybenzoate to 3, 4- dihydroxybenzoate and possess a 394 amino acid sequence that forms the characteristic two-domain PHBH fold, which houses the FAD binding domain (Van Berkel, 1995). Although the PHBH family of enzymes shares identical FAD-binding sites, their catalytic centers are markedly different giving rise to the diverse functionalities of one-component monooxygenases (Mattevi, 1998).
An exclusive characteristic of the class D and E flavoenzymes is the presence of a single gene encoding an NAD(P)H-specific reductase and FAD-dependent monooxygenase on two separate polypeptide chains, which do not need not directly interact with each other for the FAD-dependent hydroxylation phenolic compounds (Van den
Heuvel, 2004). Of these two-component enzymes in class D, the 4-hydroxyphenylacetate
3-monooxygenases from the Pseudomonasputida strain are the most well understood (Van
Berkel, 2006). These flavoenzymes convert 4-hydroxyphenylacetate to 3, 4 - dihydro- phenylacetate by means of an overall reaction that uses an HpaB oxygenase component to introduce the hydroxyl group and an HpaC reductase to supply reduced flavin needed for catalysis (Kim, 2007). Crystal structure analysis of the HpaC reductase from Thermus 18
thermophilus HB8 determined that FAD has a much lower binding affinity when compared to the PheA2 reductase component from Bacillus thermoglucosidasius A 7, which draws attention to the known structural differences in the region involved in binding the AMP moiety of FAD (Kim, 2007).
1.5 Catalytic mechanism of styrene monooxygenases
A
Figure 6: Oxidative and Reductive Half-Reactions of (A) One- and (B) Two-Component Monooxygenases. Enzyme bound flavin is reduced by NAD(P)H in the reductive half reaction (blue), and the reduced flavin interacts with oxygen and the substrate (S) to form the oxidized product (P) in the oxidative half-reaction (purple). (A) Single-component mono-oxygenases oxidize and reduce flavin in the same active site. (B) Two-component enzymes, oxidized flavin (Flox) and reduced flavin (Fired) are transferred between the reductase (El) and oxygenase (E2). Red color depicts reaction path producing reactive oxygen species (Sucharitakul, 2014).
Non-enzymatic reduction of free flavin by other reduced flavin nucleotides is generally a slow process that has been facilitated by the evolution of a flavin reductase component capable of reducing riboflavin, FMN, or FAD by NADH or NAD(P)H (Van 19
Berkel, 2006). Reduced flavin plays a vital role as a redox mediator in the metabolism of aromatic substrates in two-component flavoprotein systems like styrene monooxygenase from Pseudomonas sp. VLB 120 bacteria (Morrison, 2013). Although, the reaction of reduced flavin with oxygen is complex, these redox enzymes utilize the electron rich flavin intermediate to form a semi-stable C4a-hydro peroxyflavin adduct, which aids in the splitting of the oxygen-oxygen bond and incorporation of a single oxygen atom into an organic substrate to catalyze hydroxylation and epoxidation reactions (Van Berkel, 2006).
Recent crystallographic data on the SMOB reductase component highlights the importance of the N-terminus in the regulation of flavin reduction and elucidated the binding environment of the isoalloxazine ring structure (Morrison, 2013). The differences in the general catalytic mechanisms of one- and two-component monooxygenases are shown above in Figure 6; 6B depicts the oxidative and reductive half-reactions that will be elucidated in the next section. Wild-type styrene monooxygenase (SMO) catalysis is facilitated by a set of FAD-binding equilibria that supports the efficient exchange of flavin between the SMOB and SMOA components, and studies have proven that SMOA has a 5- fold higher affinity for reduced FAD than SMOB (Morrison, 2013). The catalytic mechanism of wild-type SMOB is similar to the HpaC component of 4- hydroxyphenylacetate 3-monooxygenase, and also displays decreased reduced FAD- binding affinity (Morrison, 2013; Kim, 2007). Furthermore, the reductase has a 1000-fold higher affinity for oxidized flavin than SMOA (SMOB 1,2|uM and SMOA 1,6mM), which 20
explains how SMO is able to prevent the production of reactive oxygen species allowing the successful return of FAD to SMOB after styrene epoxidation (Morrison, 2013).
Steady-state kinetic experiments indicate that at low FAD concentrations apo-
SMOB catalyzes the reduction of FAD using an ordered sequential mechanism with
NADH as the leading substrate (Kantz, 2005, Otto, 2004). The fluorescent C4a- hydroperoxyflavin intermediate forms at the rapid rate of ~1000 s'1, with the kinetics of the epoxidation reaction being rate limited by the preceding SMOB hydride-transfer reaction
(Morrison, 2013). In the absence of styrene, SMOA accepts reduced flavin from the SMOB active site through a direct inter-protein interaction with the SMOA-hydroperoxyflavin and apo-SMOB (Morrison, 2013). However, in the presence of styrene the peroxy intermediate reacts with oxygen to allow the regeneration of oxidized FAD, a cycle that proceeds due
SMOBs high affinity for oxidized flavin (Morrison, 2013). Furthermore, a putative SMOB-
SMOA interaction may aid in altering the FAD-bound SMOB conformation from a less reactive, sequestered (S state), to an exposed (T state) to expedite the hydride and flavin transfer reactions (Morrison, 2013).
1.5.1 SMO oxidative- and reductive half-reactions
During the reductive half-reaction, the SMOB reductase component catalyzes the reduction of FAD through an important hydride transfer reaction from NADH, which is rate-limiting in the epoxidation of styrene to form SO (Montersino, 2011) and proceeds at
50 s'1 (Morrison, 2013). Following the hydride transfer reaction, reduced FAD leaves the
SMOB active site to bind SMOA with high affinity where it quickly reacts with O2 to form 21
a stable C4a-hydroperoxy intermediate during the oxidative half-reaction of two- component monooxygenases shown in Figure 6B. Once SMOB has transferred its reduced
FAD to SMOA it is shut-off and undoable to re-oxidize NADH allowing the C4a- hydroperoxide within the SMOA active site to react with its styrene substrate (Kantz, 2011;
Ukaegbu, 2010). In the presence of styrene, the C4a-hydroperoxide intermediate can react rapidly to produce styrene oxide and a C4a-hydroxyflavin, which then eliminates water to regenerate oxidized FAD.
Recent discoveries related to the mechanistic models of how flavin-dependent monooxygenases control the reaction with oxygen has highlighted important features of the isoalloxazine ring system and the FAD-binding environment (Chaiyen, 2012). Present research on the catalytic mechanisms on the reductive half-reaction of two-component monooxygenases suggest that depending on the enzyme structure or aromatic substrate, the overall catalytic cycle can occur through an ordered sequential mechanism or a ‘ping-pong’ double displacement mechanism (Mattevi, 2006). Increased binding affinity of the FAD prosthetic group, Kd = lOnM, in the PheA2 reductase component of Bacillus thermoglucosidasius A7 has been implicated in its preference for the double-displacement mechanism (Van den Heuvel, 2004). Crystallographic results indicate that exogenous FAD binds in the NADH pocket following release of NAD+ and retains the ability to bind one
FAD cofactor and one FAD substrate (Van den Heuvel, 2004), in contrast to the catalytic mechanism of the well-studied two component flavoenzyme, 4-hydroxyphenylacetate 3- monooxygenase (Kim, 2007). In both enzymes the reduced flavin reacts with molecular 22
oxygen in the active site of the monooxygenase to generate fluorescent peroxide intermediate that subsequently oxidizes the styrene substrate. Moreover because these proteins carry out their reductive and oxidative half-reactions in two separate polypeptide chains, their direct interaction is may or may not be mandatory for efficient reduction and epoxidation to occur (Van den Huevel, 2004).
1.5.2 SMO regulation andflavin-exchange reactions
Organic flavin cofactors represent a family of versatile redox molecules in the chemistry of life, with a large proportion of eukaryotic and prokaryotic genomes encoding for FAD and FMN (Mattevi, 2006). The tricyclic isoalloxazine ring system is a signature flavin feature; its fused hydrophobic dimethylbenzene rings form an amphipathic molecule with a hydrophilic pyrimidine ring, possessing a two-electron reduction potential of about
2200 mV (Fraajie, 2000). Equilibrium-binding studies indicate that the SMOA epoxidase binds reduced flavin ~13 times tighter than the reductase, which facilitates its reaction with molecular oxygen and the formation of the activated C4a-hydroperoxyflavin intermediate 23
(Morrison, 2013); Figure 7 illustrates the changes in the isoalloxazine ring structure during the catalytic cycle of flavin-dependent monooxygenases.
C4eHKydroperaxyflavin
Figure 7: Isoalloxazine Catalytic Cycle of Flavin-Dependent Monooxygenases. During the resting state the reduced nicotinamide cofactor NAD(P)H binds to the flavoenzyme and transfers a hydride to the isoalloxazine ring (1). Next (2), reduced flavin reacts with molecular oxygen to produce the catalytically active C4a-hydro- peroxyflavin intermediate, which (3) mono-oxidizes the aromatic substrate. (4) The resulting C4a-hydroxyflavin dehydrates to reform oxidized flavin (Hotlmann, 2014).
In specific terms, the initial step in the reaction with O2 is the transfer of one electron from the reduced flavin to O2 to generate a caged radical pair (superoxide anion and flavin semiquinone) that results in the production of a one electron reduced flavin
(Massey, 1994). This initiation step is needed to overcome the spin-inversion barrier due to differences in the singlet-state of reduced flavin and the triplet-state of molecular oxygen
(Chaiyen, 2012). The regulation of FAD-binding was proposed to be dependent on the 24
SMOB equilibrium between the unreactive (S state) and a reactive transfer (T state)
(Morrison, 2013). During the hydride-transfer reaction, subsequent dissociation of NAD+ transiently occupies the T state thus promoting the transfer of reduced flavin to SMOA
(Morrison, 2013). The proposed S/T state model takes the binding of the NADH pyrimidine dinucleotide to SMOB into account, implying that this step causes the shift in equilibrium to from the T to S state that directly affects the epoxidation kinetics and structure of SMOB
(Morrison, 2013).
In the presence of an alternate electron acceptor, cytochrome c, flavin reduction also proceeds at a rate of ~50 s'1, equivalent to the wild-type hydride-transfer reaction, which confirmed that the rate of reduced flavin dissociation from SMOB is much faster than 50 s'1, and is rate limited by the preceding hydride-transfer and oxidized pyrimidine nucleotide dissociation reactions (Morrison, 2013). Although wild-type SMOB favors an ordered sequential mechanism during the reductive-half reaction involving flavin, new studies continue to implicate the N-terminal region in modulation of the flavin binding environment and therefore the overall SMO catalytic mechanism (Morrison, 2013). 25
1.6 Styrene monooxygenase structure
Figure 8: N-terminally Tagged SMOB. The location of FAD binding is shown in yellow and the location of the 6-histidine N-terminal tag is depicted using a 20-amino acid space filling model.
Styrene monooxygenase catalyzes the epoxidation of styrene through the expression of StyB, an NADH-specific reductase and StyA, an FAD-dependent monooxygenase from Pseudomonas sp. bacteria. The presence of the many similar and differential structural relationships of two-component flavoenzymes raises the question of how their physical characteristics impact their mechanistic consequences (Sucharitakul,
2014). 26
1.6.1 SMOB: Styrene monooxygenase B, reductase features
An N-
fcQG* terminally tagged
t. - "vSv^ version off SMOBcMo r was ^ e \\ ' » IJr H ' v - \ViAjX used for x-ray
I * I 1 x ■ '% ' it i i Yj \ i '>«vf crystallography, the
crystal structure of the
20kDA SMOB unit Figure 9: Overall Structure of N-terminally Tagged Styrene Monooxygenase A. (A) NSMOA monomer. was solved with a 2.2 Domain A is colored blue and domain B green. NSMOA secondary structure. (B) NSMOA dimer colored as in A resolution. It was panel A. (Ukaegbu, 2010). determined that the reductase is a homodimeric protein with a two-fold center of symmetry with the active site serving as the molecular interface between the two subunits (Morrison, 2013).
The SMOB FAD-binding fold was determined to be similar to that of the HpaC and
PheA2 reductases, homologous proteins from known two-component (Morrison, 2013).
Due to the highly disordered nature ofthe N-terminal tag, the electron density in this region was not able to be accurately resolved, however given the position of the first observable residue on the N-terminus, this region is most likely located near the FAD/NADH binding pocket (Morrison, 2013). As a consequence of crystal structure packing, each SMOB 27
monomer also binds an FAD molecule at different sites of the same face of the protein demonstrated in Figure 8 (Morrison, 2013).
1.6.2 SMOA: Styrene monooxygenase A, epoxidase features
The crystal structure of an N-terminally histidine-tagged SMOA was solved at a
2.3A resolution, and indicated that the enzyme exists as a 46 kDA homodimer with two distinct domains shown in Figure 9 (Ukaegbu, 2010). The overall architecture of SMOA is homologous to the single-component PHBH enzyme, even though the secondary structure is significantly altered (Ukaegbu, 2010). Physical comparisons show the presence of a large cavity that forms the FAD-binding site near the surface and a styrene binding site towards the base of the monooxygenase (Ukaegbu, 2010). Redox experiments confirmed the presence tightly coupled system becasue reduced FAD binds apo-SMOA
-8000 times tighter than oxidized FAD (Ukaegbu, 2010). In light of these structural and mechanistic features, increased concentrations of either component also augment the FAD- binding affinity by ~ 60-fold (Ukaegbu, 2010). Furthermore, the styrene epoxidation reaction is rate limited by the reductive half-reaction involving the flavin-transfer and oxygen binding, which emphasizes the importance of apo-SMOA as the catalytic resting state of the oxidative half-reaction (Ukaegbu, 2010).
1.7 Naturally-occurring and artificially engineered fusion proteins
The results introduced above support the idea that the flavin-transfer mechanism occurs through a functional SMOB-SMOA reaction complex the free diffusion of excess reduced FAD, an interaction that has been shown to stabilize the reductase, allow it to 28
receive re-oxidized flavin from SMOB and prevent protein aggregation (Morrison, 2013).
The array of flavin exchange mechanism was recently expanded by the discovery of a new class of styrene monooxygenase fusion proteins from Rhodococcus opacus 1CP (Tischler,
2009). This species encodes a self-sufficient StyAl monooxygenase and a fused StyA2B,
NADH-flavin oxidoreductase component on a single polypeptide chain (Tischler, 2009).
In general, this novel system proceeds using flavin intermediates similar to the two- component styrene monooxygenase from Pseudomonas sp. , but catalyzes the synthesis of styrene at a considerably slower rate than the P. putida S12 group (Tischler, 2010).
Regulation of the StyAl/StyA2B system is may be similar to the typical StyA/StyB enzyme from P. putida', experiments using equimolar ratios of each component provided the highest monooxygenase activity, highlighting the possibility of a transient protein- protein complex also (Tischler, 2010). Although the independent StyAl epoxidase is active when reduced flavin is supplied by PheA2 or SMOB, the high rate of uncoupling produces
A StyAiStyB B. StyAZB C. StyAl I StyA2B siymrt* o*sd*
Figure 10: Genetic Organization and Mechanism of SMO Systems. (A) Typical mechanism of StyA/StyB from Pseudomonas sp. VLB120 (B) Mechanism of the StyA2B fused oxidoreductase from R. opacus 1CP. (C) Putative complex and mechanism of StyA2B /StyAl monooxygenase. Excess reduced flavin generated by the StyA2B reductase is utilized by StyAl, increasing styrene’s oxygenating capacity. Dashed arrows indicate uncoupling-based FADH2 auto-oxidation leading to the formation of hydrogen peroxide (Tischler, 2010). 29
reactive oxygen species because the epoxidation efficiency is highly dependent on the type
of reductase used (Figure 10) (Tischler, 2010). In addition to the external determinants of the StyA 1/StyA2B system, the StyA2B monooxygenase unit of was found to have little
oxygenating ability functions primarily as a generator of reduced flavin (Tischler, 2010),
questioning the evolutionary significance of this novel multifunctional flavoprotein
monooxygenase.
Both the mechanistic and structural studies of SMO have demonstrated roles for
both redox-linked coenzyme- binding equilibria of SMOA-SMOB complexes in the
regulation of styrene oxide synthesis (Kantz, 2011; Morrison, 2013). In light of the StyA2B
discovery, engineered fusion proteins have been shown to also display similarities in the
reductive-half reaction (Tucker, Unpublished). Structurally, the naturally-occurring
styrene monooxygenase fusion proteins, StyA2B, occur with a reductase domain linked to
the C-terminus of the epoxidase domain (Tischler, 2013). Furthermore, comprehensive
studies on fusion proteins have been limited due to SMOs two-component nature,
disadvantages in recombinant expression, enzyme purification, and reduced efficiency in
the flavin-transfer reaction (Tischler, 2012).
Preliminary data suggests that it may be possible to create single-component
styrene monooxygenases by artificially fusing the Pseudomonas derived StyB reductase
and epoxidase components into a single polypeptide (Tischler, 2010). Engineering studies
focused on the characterization of StyALIB and StyAL2B styrene monooxygenases (kDA
65), which join the reductase and epoxidase components with unique linker peptides 30
(Tischler, 2010). StyAL2B was expressed with a longer peptide linkage and of the two is more stable, underscoring the impact of adding a component to the N-terminus of the reductase, which may affect C4a-hydroperoxyflavin intermediate stability and the reductive kinetic mechanism (Tucker, Unpublished).
In manufacturing efficient monooxygenases as biocatalysts, it is important to evaluate the structural and mechanistic importance of the N-terminal linkage of the reductase to C-terminus of the epoxidase (Tischler, 2010). Biotechnology and future research on two-component monooxygenase systems are focused on optimizing the FAD transfer-reaction to minimize the production of hydrogen peroxide and accelerating these the epoxidation of aromatic substrates, which is expected to increase catalytic efficiency and the value of StyA2B and similar reductases as biocatalysts (Bommarius, 2011; Lin,
2012). While the generation of new bioremediation strategies geared towards managing xenobiotic pollution has flourished through genetic engineering, new research is still needed to elucidate the genetic stability of heterologously expressed genes, and application of engineered fusion proteins in medicinal chemistry (Holtman, 2014).
1.8 Putative complexes
The discovery of naturally occurring fusion proteins from Rhodococcus opacus
1CP, which integrates the reductase and epoxidase function on a single polypeptide introduced new opportunities for studying the bioengineering capabilities of styrene monooxygenase enzymes (Tischler, 2012). Additionally, artificially engineered fusion proteins from Pseudomonas sp. that link the epoxidase subunit to the N-terminus of the 31
reductase, have been shown to possess differential stability, epoxidation capabilities, and utilize a completely different mechanism for flavin reduction during steady-state catalysis
(Tischler, 2013).
Based on previous data and preliminary results we propose that research should be aimed at characterizing the engineered N-terminally tagged SMOB reductase and evaluating the impact that the addition of the 20-amino acid moiety has on: FAD binding affinity, the steady-state catalytic mechanism, and the rate-limiting hydride transfer step.
Previous data on structural features that would influence a putative StyB/StyA complex stressed the importance of the N-terminus in the regulation of flavin reduction and elucidated the binding environment of the isoalloxazine ring structure within the PHBH fold (Mattevi, 1998). Figure 11 depicts the putative physical location of the N-terminal tag and suggests that the presence of this moiety may directly affect the SMOB-SMOA flavin- transfer reaction and explain the change in catalytic mechanism between wild-type SMOB and both the naturally-occurring and engineered fusion proteins, which link the N-terminus of StyB to the C-terminus ofthe StyA subunits (Tischler, 2010; Tischler, 2013; Morrison,
2013). Figure 11: (A) Complex Formation of the Wild-Type Styrene Monooxygenase Reductase and Epoxidase Components during the FAD-Exchange Reaction of the native enzyme. (B) Histidine tag interferes withNSMOB to NSMOA FAD-Exchange.
1.9 Transition to thesis research
Despite the fact that much of the styrene side-chain oxidation pathway has been elucidated at the biochemical and genetic level, little attention has been focused on studying the physiological factors affecting the regulation of the pathway (Van Berkel,
2006). This kind of information is invaluable in expediting the use of styrene degradation enzymes in bioremediation and present a new frontier in advancing the manipulation of metabolic pathways for biotransformation applications for the production of optically pure chemicals with broad chemical reactivities (O’leary, 2001). Significant aspects of styrene- induced transcriptional regulation that are presently unresolved making it unclear if manipulation of StyB reductase components to enhance purification yield and epoxidation 33
efficiency are truly promising with emphasis on the circumstances that exert negative influences on catalytic mechanisms of all StyA/B system (O’leary, 2002).
In this research we evaluate the impact of an engineered 20-amino acid SMOB N- terminal 6-histidine tag on the FAD binding affinity, the steady-state catalytic mechanism, and the rate-limiting pyridine nucleotide to FAD hydride transfer and mechanism of reduced FAD transfer from SMOB to SMOA. These results have important implications in assigning function to the N-terminal structural domain of SMOB as it occurs in the two- component SMO system and provides new insight into the mechanism and engineering of styrene monooxygenase fusion proteins alike. 34
METHODS
2.1 Protein expression and purification
Styrene is both unavoidably integrated in our daily lives as significant health concern, and we have focused our research efforts on elucidating the structures and mechanisms of, styrene monooxygenase B, the entry point of styrene into the detoxification pathway. Toward this goal we have purified a wild-type and N-terminal expression system for SMOB and investigated its catalytic mechanisms, the detailed methods are presented below. Additionally, the cloning design of expression vectors and sequencing is detailed in a previous paper (Kantz, 2005), and included the DNA isolation from Pseudomonas putida SI2 bacteria with primer design based on the styA and styB sequences of Pseudomonas sp. (Kantz, 2005).
2.1.1 Native SMOB purification
In general the expression of native SMOB involves the expression of the reductase in E. coli BL21 (DE3) cells with a ~30mg/liter yield, and a similar protocol to the one described for NSMOB in section 2.1.3 (Kantz, 2005). Our SMOB experiments used a PET-29-SMOB BL21(DE3) cell pellet, which was thawed, sonicated 6 x 30 seconds, and then centrifuged at 6000 rpm x 10 minutes. The pellet was washed in 100 ml of wash buffer A (50mM TRIS pH 7.5, 0.5% Triton-X 100, lOmM EDTA), 100ml of wash buffer B (50mM TRIS, 7.5 pH, 2M urea), incubated for 10 min at 4°C, and centrifuged at 2000 x g for 30 minutes. Then 50 ml of the SMOB pellet was re-suspended 35
in 125 ml of wash buffer C (50mM Tris buffer, 10 mM DTT, 8M urea) for a total volume of 120ml and urea concentration of 4.2M. Previous studies indicated that only about 1% of the SMOB expressed is located in the soluble fraction and the best methods to recover soluble and active SMOB from the inclusion bodies is to use 8M urea containing 10 mM
DTT (Otto, 2004; Kantz, 2005; Morrison, 2013).
2.1.2 Recovery of soluble SMOB from inclusion bodies andflavin dialysis
120 ml of 8M urea-denatured SMOB was separated into three 40 ml fractions.
Each SMOB fraction was refolded in the presence of lOOpM FAD, FMN or riboflavin, and dialyzed overnight using Spectra/por membrane tubing (MWCO 6-8,000) against 1 liter of (50mM tris buffer, ImM DTT). Each flavin-refolded SMOB sample was clarified for 10 min x 15,000 rpm for a yield of 37ml, 38ml, and 34ml for FAD, FMN and riboflavin-refolded SMOB, respectively. 30 ml of each dilute sample was then concentrated by 1/3, and both were stored without glycerol at 4°C. After day 8 all samples were stored in 50% glycerol at -20°C.
2.1.3 N-Terminally tagged SMOB expression and purification
The N-terminally histidine tagged version of SMOB was expressed from the pET-
29 NSMOB vector in E. coli BL21 (DE3) cells as described in a previous protocol
(Kantz, 2005). A 6 liter N-His SMOB preparation was completed and E. coli cells were grown at 37°C in 30 pg/ml kanamycin and induced with ImM IPTG for 60 minutes. The cells were then harvested when OD 600 = 1. The pellet was stored at -80°C. Cell pellets were sonicated for 6 x 30 seconds in a buffer containing ImM PMSF, 100 nM EDTA in 36
50 ml of lOmM imidazole buffer pH 8 containing 300 mM NaCl. The resulting suspension was pelleted by high-speed centrifugation at 20,000 rpm x 30 minutes in an
SS34 rotor.
Immediately after centrifugation soluble NSMOB supernatant was recovered and loaded onto a His-Select nickel-affinity column on a BioRad BioLogic HR FPLC at a flow rate of 3mL/min (Kantz, 2011). The column was equilibrated with a pH 8 buffer containing 300 mM NaCl and 10 mM imidazole followed by a linear gradient up to 300 mM imidazole. Fractions containing NSMOB were recovered based on UV-absorbance
Appendix 1 and up brought to 100 |^M in FAD and concentrated in centriprep-10 concentrators to reduce the sample volume by 1/3. The concentrated enzyme was stored in 50% w/w glycerol at -20 °C, samples have a half-life of 2-days at 4°C, but can be stored indefinitely at -20°C in a stabilization solution of 50% glycerol.
2.2 Activity and purity assays
Both the native and N-terminally tagged purified protein was analyzed by SDS-
PAGE, the NSMOB gel is shown in Appendix 2. The SDS-PAGE results of the wild- type protein after subsequent flavin-refolding will be discussed in the following results section. We used the activity assay to determine the specific activity and total protein concentration using a Thermofisher Scientific Pierce BCA Protein assay kit with BSA standards using a method that was previously described (Kantz, 2005). 37
2.2.1 SDS -Polyacrylamide Gel Electrophoresis
An SDS-PAGE analysis was run using a 12% polyacrylamide gel to detect the amount of protein present during specific steps of the protein purification and recovery process. 20pl of each sample and molecular weight markers were dissolved in 20pl of 2x running buffer and denatured for 90 seconds. 20pl of each denatured sample was loaded onto the gel and run under a constant current of 30mA and 100-160V for 70 minutes.
2.2.2 Microplate Reader-Based BCA Assay
A plate reader based BCA assay was performed on non-concentrated samples with a Spectromaxl90. A BSA standard curve was obtained as detailed in previous experiments (Kantz, 2005). 50pl of diluted SMOB was added to 1ml of BCA reagent, and incubated at 37°C for 30 minutes. 250 pi of each sample was added to a polystyrene p-Plate and monitored at 562nm. The NSMOB BSA standard curve Appendix 3 and total protein results are shown in Appendix 4; the results for the flavin-refolded SMOB will be presented in the results section.
2.2.3 Gel filtration
Gel filtration and small scale dialysis was tested for their ability to remove the imidazole, glycerol, and free flavin from all flavin-refolded SMOB samples. All SMOB and NSMOB samples underwent gel filtration before any kinetic measurements were taken; those that were not will be noted and explained in the results section. Gel filtered samples were eluted in 20mM MOPSO buffer pH7 from a Bio-Rad® DG-6 column. 38
Alternatively due to the frequent reduction in enzyme activity after gel filtration, we experimented with small scale dialysis to remove the imidazole and excess flavin from SMOB. 500|iL samples were dialyzed using 0.1-0.5ml Slide-A-Lyzers® in 1000ml of 20mM Tris buffer pH7 for 4 hours. We also performed kinetic assays to evaluate the impact of gel filtration on enzyme activity; the importance of our findings will be presented in the results section.
2.2.4 Flavin-refolded SMOB microplate reader-based activity assay
The kinetic activity of each flavin-refolded SMOB sample was observed using a
Spectromaxl90 microplate reader; NADH absorbance was monitored at 340nm for 2 minutes. Activity assays were run using FAD, FMN, or riboflavin as the catalytic flavin for each of the flavin-refolded SMOB proteins. 195^1 of MOPSO pH 7 buffer, 5^1 of diluted protein, and 25|xl of 300pM flavin were added to each well. 25(xl of ImM NADH was added and mixed thoroughly to bring the total volume to 250^1 with a final catalytic flavin concentration of 30|aM.
2.2.5 NSMOB microplate reader-based activity assay
The kinetic activity of each flavin-refolded SMOB sample was observed using a
Spectromaxl90 microplate reader; NADH absorbance was monitored at 340nm for 2 minutes. Activity assays were run using FAD, FMN, or riboflavin as the catalytic flavin for each of the flavin-refolded SMOB proteins. 195pl of MOPSO pH 7 buffer, 5pl of diluted protein, and 25pl of 300|iM flavin were added to each well. 25pl of ImM NADH was added and mixed thoroughly to bring the total volume to 250pl with a final catalytic 39
flavin concentration of 30pM. Due to the differential protein concentrations determined by BSA analysis we diluted each protein to account for the differences in activity based on the greater fraction of active and refolded SMOB: FAD 50X, FMN 10X and riboflavin
5X.
2.3 Time-dependent activity studies
Due to the varied kinetic activity that we observed from day to day experiments we decided to monitor the effects that time, gel fdtration, temperature, and glycerol had on the overall reductase stability. The experimental methods for these studies are detailed below.
2.3.1 Flavin-refolded SMOB half-life studies
The kinetic activity of each flavin-refolded SMOB sample was observed over a 9 day period using a Spectromaxl90 microplate reader to observe the effect that time and glycerol have on the kinetic activity of SMOB. NADH absorbance was monitored at
340nm for 2 minutes. Activity assays were run using FAD, FMN, or riboflavin as the catalytic flavin for each of the flavin-refolded SMOB proteins. 195jj.1 of MOPSO pH 7 buffer, 5pl of diluted protein, and 25pl of 300pM flavin were added to each well. 25pl of
ImM NADH was added and mixed thoroughly to bring the total volume to 250pl with a final catalytic flavin concentration of 30pM. The A340/sec values were converted to
A340/min and multiplied by ~500-fold to account for the dilution during plate reader assays. Equation 1 was used to fit the velocity data with vO being the rate taken previously and vl being the rate observed for that given day. 40
r- In 2 -] ------r r ,/ L 2 J Equation 1: Velocity
II 1 — e rate equation for time H* O dependence studies.
2.3.2 Absorbance and fluorescence spectra offlavin-refolded SMOB
Concentrated, FAD-refolded SMOB samples were gel filtered for spectral analysis and fluorescence analysis was completed using a Horiba-Jorbin-Yvonne
Fluorimeter. Excitation and emission scans were obtained for FAD-reconstituted SMOB.
A fluorescence monitored titration experiment was performed using 500nM of FAD- reconstituted SMOB stored at 4°C; the relative emission at 563nm was observed.
2.4 Steady-state reaction mechanism of NSMOB
Steady-state kinetic data was recorded using a Molecular Devices SpectraMax
190 Multi-Mode Microplate Reader with SoftmaxPro software. A 64-well polystyrene plate with a pathlength of 0.755 cm was used. The kinetic activity of purified N- terminally histidine-tagged SMOB was evaluated by analysis of NADH consumption at
A340nm, and three trials were recorded for each concentration of FAD and NADH. All samples and reagents, aside from the enzyme, were kept in a water bath at 30°C prior to the experiments, and the plate reader was set at 30°C. NSMOB enzyme was kept on ice and diluted by 15X into 20mM pH 7 MOPSO reaction buffer, which was determined to give the best results based on a simple protein concentration assay. 5(il of diluted 41
NSMOB was then added to 25fil of catalytic FAD, which varied from 2 |xl to 50 jj.1, and
195|al of MOPSO buffer. All reactions were initiated by adding 25^1 ofNADFl at varying concentrations from 2 fx! to 50 (il for a for a total reaction volume of 250 pL and mixed thoroughly. NADH absorbance was monitored at 340nm for 2 minutes and the initial rate vales were recorded in the first 10 to 20 seconds of data collection. The use of the initial rates is important because the first 5-10 seconds represents the oxidation of NADH to
NAD+, and does not reflect the reduction of FAD due to its rapid reoxidation on the presence of oxygen (Kantz, 2005).
2.4.1 Determination of NSMOB catalytic mechanism
3 replicate reactions were analyzed for each experimental condition and the reaction rates were compiled to create a single data array and fit globally along the substrate concentration axis to describe the reaction (Kantz, 2005). The apparent Vmax and Km values were determined from the fit using KaleidaGraph and are given by the equations in Equation 2.
Vmaiapp Equations Equation 2: Kinetic ,. [NADH] [FAD] r,W l/°»' _____ Fitting Equations. V max “ ~ K m™ + (NADH] “ KT + [**£>] apparent and Km apparent equations. The apparent Kmapp Equations Km and V m a x parameters were used in GraphPad k T wadh] K ^ \ fad] Prism software global- KT h +\n a d h ] A m k ™ + [f a d ) fitting analysis.
v v%[ f a d \ K°pp + [FAD] 42
Microplate-reader based A/340 kinetic data was the converted to velocity measurements and data points analyzed were weighted based on standard deviation. Once the data was sufficiently transformed with the best representative concentration of NADH or FAD axis data, we compared the observed NSMOB values to the native enzyme. The velocity (v), Km and Vmax apparent, are a function of [FAD] and [NADH]. The steady- state rate equation for the ordered BiBi sequential mechanism gives the definition of the
Km apparent parameter on the left in Equation 3, with the difference in mechanism being highlighted by the Ks value. Km apparent parameter for the double displacement mechanism in Equation 4.
Figure 3: Ordered Sequential Mechanism. The velocity equation is a function of A= [NADH] and B= [FAD], and the Km apparent equation.
V
+ [NADH]
Equation 4: Double-Displacement Mechanism. The velocity is a function of [FAD] and [NADH], 43
KaleidaGraph 4.0 was used to determine the best fit curves for the steady-state experiments and the rate vs. [NADH], Km of NADH, Km of FAD and Vmax ofNSMOB were calculated. GraphPad Prism 4.0 software was used for the global fitting of the steady-state data and the compare models feature selected the best mechanistic model between the Bibi sequential and double displacement mechanisms (Kantz, 2005;
Motulsky, 2004).
2.5 Estimation of equilibrium dissociation constants by fluorescence monitored titrations
Fluorescence analysis was conducted with a Horiba Jobin Yvon Fluorolog-3 spectrofluorometer to estimate the FAD-binding affinity, Kd of N-terminally tagged
SMOB. Enzyme was exchanged into 20mM MOPSO buffer, pH 7.0 by gel filtration to remove excess FAD and diluted to a concentration of 500 nm in a 3 ml quartz cuvette containing 1.5 ml of buffer. Fluorescence excitation at 456 nm and emission spectra at
563 nm were recorded with 1 nm spectral resolution at 4°C while stirring the sample with a 5 mm Teflon-coated stirrer bar. The data were fit using the quadratic in Equation 5. 44
F = EdFADltotal +
K PP [FAD]** + [SMOA]total - (K“pr>+{FAD]tow! + jS M O B ]^ y -4[FAD]totai[SMOA]R J (£2 -ei)
Equation 5: FAD-Binding Constant Quadratic Equation.
The interaction between oxidized FAD and NSMOB was observed at equilibrium as the increase in steady-state fluorescence due to SMOB binding flavin. Oxidized FAD is known to bind tightly to native SMOB and the increases in fluorescence were used to compute the apparent binding constant of FAD in the presence of SMOB, with £1 and £2 being the molar extinction coefficients for the fluorescence of free FAD and bound FAD.
Fits passing through the titration data for native and N-terminally tagged SMOB will be discussed in the results section and standardization of FAD bound was used to account for the amount of FAD already existing is solution after gel filtration.
2.6 Pre-Steady State fluorescence and absorbance kinetic analysis
Single-turnover kinetic data were recorded using an Applied Photophysics SX-17 stopped-flow instrument equipped with absorbance and fluorescence photomultiplier tubes as previously described (Kantz, 2005). NSMOB enzyme was exchanged into lOOpM NaCl and 50pM Tris buffer at pH 7.0, to preserve the integrity of the enzyme and remove excess FAD. Previous studies indicate that NSMOB prefers to be in an environment of higher ionic strength during gel-filtration, and kinetic assays proved this 45
when compared to samples gel-filtered in MOPSO buffer at pH 7.0. NSMOB sample were diluted to a concentrations of 2 - 3 |xM; all kinetic studies were performed at 15°C.
The stopped-flow instrument was controlled by SpectraSuite software and triggered externally by a stop syringe. The stopping syringe allows very small volumes to be injected into the cell, and the flow time is designated as the time it takes for the solution to fill up the cell before mixing with a dead time of about 2 ms.
Stopped flow data was used to investigate the single turnover reactions: absorbance spectroscopy at 340nm for observing the steady-state oxidation of NADH and reduction of FAD at 450nm, and fluorescence excitation of FAD at 450nm and emission at 520nm. Logarithmic time averaging reduced the size of the data set and amplified the low signal readings and will be discussed in the results section. All corresponding kinetic spectra were fit with exponential equations using KaleidaGraph 4.0 software to give the rate constants. 46
RESULTS
3.1. Flavin-refolded SMOB experiments with riboflavin, FMN and FAD
SMO is a two-component flavoenzyme composed of NADH-specific reductase
(SMOB) and FAD-dependent monooxygenase (SMOA) that catalyzes the enantioselective epoxidation of styrene to (S^-styrene oxide. In catalysis, SMOA and
SMOB share a single FAD using a flavin-exchange mechanism. Flavin reduced by
SMOB is transferred from the active site of SMOB to apo-SMOA where it binds tightly and catalyzes the activation of molecular oxygen in the styrene epoxidation reaction.
However, the amount of the folded reductase enzyme is much higher when a flavin is present and activity and binding assays have shown that the optimal yield of active
SMOB is reached when it is tightly bound to oxidized FAD (Kantz, 2005).
Over expression of the
wild-type SMOB produces a / / -r#’t&m high yield of the reductase in the
form of insoluble inclusion
bodies, which can be denatured
Figure 12: Flavin-mediated refolding of SMOB in urea and refolded using in 8M urea. dialysis illustrated in Figure 12.
Using the protocols detailed within the methods section, we sought to determine if the characteristic flavin isoalloxazine ring is the primary factor that determines the specificity 47
of SMOB refolding or if the AMP moiety also mediates the necessary interactions for optimal yield and protein activity.
Our goal was to determine whether the specificity of refolding and delivery of reduced flavin to the FAD-specific SMOA epoxidase will be affected by using FMN or riboflavin. In the presence of FAD the yield of active soluble protein is much higher than refolding SMOB without FAD (Kantz, 2005). In this section we evaluate the hypothesis that FMN, riboflavin, and FAD, may each nucleate the folding of SMOB and stabilize it in a catalytically active form, the differences in flavin structure are shown in Figure 13.
Figure 13: Flavin Structure. Riboflavin, Flavin Mono nucleotide (FMN), and Flavin Adenine Dinucleotide (FAD).
First we examined our hypothesis by separately purifying the SMOB component and then using small scale dialysis to refold the reductase in an excess of FAD, FMN, and riboflavin. Next, total protein assays were completed to assess enzyme concentration and fluorescence titrations were implemented to investigate flavin binding specificity. Upon production of reagent quantities of FAD, FMN and riboflavin bound SMOB, we additionally hypothesized that in the presence of these non wild-type flavins the SMO system may engage in an indirect transfer of electrons to SMOA. Studies have shown that 48
the FAD-dependent epoxidase cannot utilize reduced FMN or riboflavin as a source of electrons so there may be an intermediate shuttling of flavin electrons to free FAD, which
SMOA can then employ to generate styrene oxide. Our preliminary conclusions predicted that the isoalloxazine ring works jointly with the AMP substituent present and both are important factors in SMOB refolding stability and protein yield.
Although we did not get the opportunity to perform single-turnover experiments with the FMN and riboflavin flavin-refolded SMOB, previous studies confirm that N- terminally tagged SMOA cannot use FMN or riboflavin as sources of electrons for the epoxidation of stryrene (Kantz, 2011). We conclude that the AMP moiety of FAD play a very large role in the flavin binding of apo-SMOA. With reduced flavin interacting with apo-SMOA in the presence of free oxidized FAD through the use of an indirect flavin- flavin electron transfer. The purity and stability of all flavin-refolded SMOB in the presence of FAD, FMN, and riboflavin are documented by SDS-PAGE and half-life studies presented in sections 3.1 and 3.2, respectively. The total yield and specific activity of each preparation is summarized in Figure 15 and Table 1.
3.1.1 SDS-Polyacrylamide Gel Electrophoresis Purification
The purification and stability of SMOB refolded in the presence of FAD, FMN, and Riboflavin are documented by SDS-PAGE in Figure 14. Figure 14: Flavin-refolded SMOB SDS-PAGE purity analysis. Lane 1, Dialysate from Riboflavin-SMOB; Lane 2, Post-clarification pellet from Riboflavin-SMOB; Lane 3, Wash buffer A supernatant; Lane 4, Wash buffer B supernatant; Lane 5, Molecular Weight Standards; Lane 6, Wash buffer A supernatant; Lane 7, FAD- refolded SMOB; Lane 8, FMN-refolded SMOB; Lane 9, Riboflavin-refolded SMOB.
All of the flavin-refolded samples showed the characteristic band between 21.5 and 31 kDa. And the washing steps were done to remove the impurities from each sample and Lane 2 represents the post-clarification pellet that displays no significant protein band indicating that most of the protein was not lost during the riboflavin-SMOB washing steps. 50
3.1.2 Total protein and specific activity
Collectively the results discussed above indicate that SMOB is most effectively refolded by FAD and FMN is an order of magnitude less effective than FAD, with riboflavin being an order of magnitude less effective than FMN in refolding SMOB into catalytically active protein.
Total Protein Yield
140
12:0
100 I c SO 1 CL 60 I 40 20
0 FAD FMN Riboflavin Flavin
Flavin-Refolded Total Protein Total Units of Activi ty Specific Activity SMOB [mg) (pmole/rain) (mU/mg) FAD 124 + 25 1.15 9.2 FMN 110 ± 6 0.096 0.77 Riboflavin 115 ±17 0.0083 0.072
Figure 15 and Table 1: Graph of Flavin-Refolded SMOB and table of Total Protein and Specific Activity Results. FAD (red), FMN (blue), and riboflavin (green). 3.1.3 Activity Assays offlavin-refolded SMOB in FAD, FMN or riboflavin
Time (min)
Figure 16: A; Flavin-Refolded Kinetic Activity Assay, (red) FAD-SMOB, (blue) FMN-SMOB, (green) Riboflavin-SMOB.
Figure 17: Flavin-refolded SMOB Kinetic Activity Graphs. (A) Flavin-refolded SMOB with FAD as the catalytic flavin. (B) Flavin-refolded SMOB with riboflavin and FMN as the catalytic flavin. All activity data used to make the graphs was dilution corrected. 52
Collectively the above results indicate that SMOB is most effectively refolded by
FAD, with FMN being an order of magnitude less effective than FAD, and riboflavin an order of magnitude less effective than FMN in refolding SMOB into catalytically active protein.
3.2 Flavin-refolded SMOB time dependence studies
The relative activity and flavin specificity of refolded SMOB were evaluated above in section 3.1. Next we will present the implications ofthe AMP moiety in SMOB enzyme half-life and stability using absorbance and fluorescence spectroscopy.
3.2.1 Half-life of SMOB refolded in the presence of FAD, FMN, and riboflavin
Figure 18: Half-Life of SMOB in the Presence of FAD, FMN, and Riboflavin at 4°C. FAD (red), FMN (green), and riboflavin (blue).
I Li I______I______J 0 2 4 6 8 10 Time at 4°C (Days) 53
At 4°C, SMOB reconstituted with riboflavin has a significantly longer half-life than the FMN and FAD reconstituted proteins. These results were unexpected because the relative yields and corresponding specific activity measurements showed the reduction rate and refolding stability to prefer FAD, FMN and then riboflavin, respectively. In summary these results establish that FAD is not an absolute requirement for catalysis, as each of the flavin-refolded preparations is catalytically active when provided with only the flavin with which it was refolded as shown in Figure 17. In addition, the proteins refolded with riboflavin and FMN show similar catalytic flavin- specificity to the reductase refolded with FAD (Otto, 2004).
3.2.2 Absorbance and fluorescence spectra of SMOB
A. B.
Wavelength Figure 19: Absorbance Flavin Reconstituted With FAD at -20°C and 4°C. A) Absorbance spectrum of SMOB recovered and immediately stored at -20°C. B) Absorbance spectrum of SMOB stored at 4°C for 8-days. 54 A comparison of SMOB-(FAD) absorbance and fluorescence spectra recorded after storage at -20°C or 4°C is shown below in Figure 19 and 18. These experiments were performed because of the significant variance in activity and stability that we witness based on the temperature and composition of the sample (with or without glycerol) over time. Panel A in Figure 19 represents a typical FAD-SMOB absorbance spectra with the flavin peak being prominent at -450 nm. However, Panel B shows a significantly blue shifted FAD peak at 445 nm indicating that the flavin-binding environment is altered after the protein spends time at 4°C without the presence of glycerol or any added FAD for stabilization. Wavelength (nm) Figure 20: Fluorescence Spectra of Flavin Reconstituted with FAD. Fluorescence excitation and emission spectra of SMOB after storage at 4°C. The structural and mechanistic basis of the time-dependent inactivation mechanism of SMOB remains to be determined. As shown in Figure 19 the peak 55 absorbance of the SMOB electronic spectrum of shifts from 456 nm (active protein) to 445 nm (inactive protein, spectrum recorded after incubation at 4°C for 8-days). 3.2.3 Time-dependent inactivation of NSMOB Despite the apparent shift in the bound-flavin environment, we find that the equilibrium dissociation constant of FAD is not significantly changed in the two forms of the protein and is depicted in Figure 21. A. B. Figure 21: Time Dependent Estimation of Fluorescence Dissociation Constants by Fluorescence Monitored Titrations. A) Native FAD-refolded SMOB stored at -20°C. B) Native FAD-refolded SMOB stored at 4°C. Kd values of 280 nM and 310 nM (Morrison, 2013) were calculated by fitting the data with quadratic Equation 5. We conclude that SMOB shows very little flavin specificity in catalytic turnover however, both the yield of SMOB in the flavin-catalyzed protein-folding mechanism and the half-life of the folded protein are dependent on the structure of the flavin catalyst 56 (Figure 17 & 18). These findings suggest a flavin-specific folding mechanism that favors the folding of SMOB with bound FAD over FMN or riboflavin. This is FAD-specificity in the folding mechanism of SMOB may be a type of regulatory mechanism that helps modulate the activity of SMOB in response to the concentration of FAD in the cell. 3.3 NSMOB Mechanistic Studies We have engineered a version of SMOB that contains an N-terminal 6-Histidine tag, which allows the enzyme to be easily purified in a folded, active state with FAD bound in its active site (Figure 22) (Kantz, 2005). Figure 22: Ribbon Structure of N-terminally Tagged FAD-bound SMOB interacting with a Nickel Affinity Column agarose bead. O We hypothesized that the addition of this 20 amino acid moiety to the reductase component would directly affect its structure and function. This research confirms the catalytic mechanism and investigates the pre- and steady state kinetics of an N-terminally histidine-tagged version of styrene monooxygenase reductase (N-SMOB). Furthermore we believe that this change in change in the FAD-binding environment may directly impact the steady-state kinetic mechanism, which is an ordered sequential mechanism in 57 the case of the wild-type enzyme. The following sections will elucidate the steady- and pre-steady state kinetics of N-SMOB and investigate the impact of the increased FAD binding affinity on the overall reductase stability. These findings will be evaluated together in full with their implications for the hydride- and flavin-transfer reactions of the two-component SMO system. 3.3.1 Steady-state catalytic mechanism determination The primary difference between the sequential and double displacement rate equations is the product of Ks and Km (NADH) in the numerator of the sequential mechanism in Equations 3 and 4. Km (NADH) Km (FAD) Vmax Ks (NADH) pM pM pMmin"1 pM Sequential 3.0 ±0.7 6.0 ± 1.6 4.8 ±0.4 1.0'7± 1.3 Double 3.0 ±0.5 6.0 ± 1.0 4.8 ±0.3 N/A Displ. Table 2: Comparison of NSMOB Kinetic Parameters. The Km of NADH, FAD, Vmax and Ks of NADH with error margins are shown. The Ks of NADH for NSMOB is very small in the sequential mechanism while the Ks of NADH is not available for the double displacement mechanism. The previous work of Berhanegerbial Asseffa utilized stopped-flow spectroscopy to evaluate the best fitting parameters of stead-state kinetics for each mechanistic model using GraphPad Prism 4.0 software. He found that the Km of NADH, Km of FAD and V m a x for N-SMOB were determined to be 5.0 pM, 3.7 pM, and 38.0 pMs'1 at 10°C. My 58 current research expands on these studies and evaluated NSMOB at 30°C under similar reaction conditions, the parameters for Km and V m a x are shown above in Table 2 Furthermore, because the Ks value for NSMOB was found to be very close to zero, this allowed the elimination of the Ks parameter and a switch to the double displacement mechanism for NSMOB during steady-state catalysis. Double Displacement Mechanism Figure 23: Global fit NSMOB Double Displacement Mechanisms. The * 2 - 3 graphs display * 5 velocities from c i 7 reactions in which £ ’ 10 both NADH and *• 12 FAD (x-axis) J 20 concentrations in uM were varied. Lines passing through the data represents the best fits for the mechanism using GraphPad Prism software. [¥AD]!pM In addition to the calculation of the Ks parameter, a statistical model comparison feature within the GraphPad Prism 4.0 program conclusively determined that the double displacement is the preferred mechanism of flavin reduction for our N-terminally tagged enzyme (Motulsky, 2007). Complete results from the comparison of global-fits that 59 summarize the calculated kinetic parameters between the sequential and double displacement mechanisms is shown in Table 3. Sequential Double Displacement Null Hypothesis 0.3494 0.8896 (1,60) Do not reject null hypothesis Table 3: Comparison of null hypothesis of BiBi Sequential and Double-Displacement Mechanisms using Prizm® Software. Nonlinear regression analysis was used to calculate the rate vs. [NADH] and the K m ap p of NADH, K m a p p of FAD and V m axap p of NSMOB from the best-fit curves steady- state analysis Equation 2. To determine the method of NSMOB steady-state catalysis the consumption of NADH at 340nm as a function of time and varying concentrations of NADH and FAD was measured (Appendix 5). Previous data confirmed that native SMOB functions through a BiBi sequential mechanism with NADH as the leading substrate characterized by the presence of an NADH and oxidized FAD charged transfer complex during the first 5 ms of the reaction (Kantz, 2005). The research presented in this section confirms that the NSMOB reductive-half reaction operates through a ping- pong double-displacement mechanism. 60 3.4 Estimation of Equilibrium Dissociation Constants by Fluorescence Monitored Titrations The equilibrium dissociation constant of oxidized flavin for N-terminally tagged SMOB was determined using a fluorescence monitored titration experiment. As free oxidized FAD is titrated into the cuvette containing apo- NSMOB, a rapid hyperbolic increase in the fluorescence emission intensity occurs as FAD binds NSMOB, following a linear increase representing the accumulation of free FAD. 500000 | 450000 Fig 24: Fluorescence 00 titration monitoring of 8 400000 FAD to NSMOB. The c fit equation provided 1 350000 an accurate estimate £ of the equilibrium US dissociation constant e of NSMOB to be ~ 69 | 250000 nM at 4°C. o 3 200000 £ 150000 500 1000 1500 2000 2500 3000 3500 (FAD) total The oxidized FAD-binding affinity (Kd) was confirmed to be 69 nM which is nearly an order of magnitude greater than that of the wild type reductase, SMOB, Figure 24. These results are significant because it was demonstrated that the orientation of important N-terminal residues located near the FAD/NADH binding pocket and the weak electron density of certain loops in this region confers disorder and flexibility to the native SMOB enzyme (Morrison, 2013). These results support the change in catalytic mechanism that would be associated with an increased FAD binding affinity due to the addition of an N-terminal tag. Furthermore, flavin would remain tightly associated with the NSMOB component during catalysis using a double displacement mechanism rather than the bibi sequential method that the native enzyme employs. 3.5 NSMOB Single-Turnover Studies Our mechanistic and structural studies of SMOB have demonstrated roles for both redox-linked coenzyme- binding equilibria SMOA-SMOB complexes in the regulation of catalysist (Morrison, 2013). Naturally-occurring styrene monooxygenase fusion proteins (eg Sty A2B) occur with a reductase domain linked to the C-terminus of the epoxidase domain and highlight the impact of the N-terminal linkage of the reductase to C-terminus of the epoxidase (Tischler, 2009). The results in the next section further investigate the impact of the 20-amino acid N-terminal tag on single turnover hydride-transfer reaction that involves the excess oxidized flavin receiving electrons from bulk reduced flavin. A reaction that is proposed due to the increased flavin binding affinity and switch to the ping-pong mechanism for NSMOB. 62 3.5.1 NSMOB stopped-flow steady-state studies To further confirm the kinetics of NSMOB we isolated the 340 nm data, which represents the steady-state oxidation of NADH in Figure 25. We obtained relevant information about the transition from pre- to steady state catalysis in NSMOB. The data was analyzed using an exponential curve to fit the pre-steady state portion, and represents the hydride-transfer rate constant. The linear region of the fit corresponds to the steady- state rate of NADH oxidation that follows the single turnover reaction; this region is rate- limited by the FAD re-oxidation reaction that occurs in the presence of molecular oxygen. 0.82 Figure 25 and Equation 6: Steady-state NSMOB kinetic data taken on a stopped-flow device. < A340 exponential and line equation fit. 0.795 0.08 0.16 0.24 0.32 0.4 0.48 0.56 0.64 Time/sec 4340(f) = 4340 exp (~kt} vt + co n sta n t 63 3.5.2 NSMOB stopped-flow hydride-transfer kinetics Time/sec Figure 26: Time averaged single-turnover NSMOB stopped-flow data. Single turnover, stopped-flow experiments examined the pre-steady state kinetics of N- SMOB. (Blue) represents the oxidation of NADH at 340nm, (Green) fluorescence excitation at 450nm and emission at 520nm, and (Red) reduction of FAD at 450nm. Logarithmic time averaging was done to alleviate issues regarding low protein concentration and small changes in signal. Fits through the data gave an average hydride rate transfer constant k = value of 48 s'1. Stopped-flow analysis was used to study the kinetics of the single-turnover hydride transfer reaction of the NSMOB reductase using fluorescence measurements. For these particular experiments the concentrations of NSMOB and oxidized FAD were rapidly mixed with 50 pM NADH and 20 pM FAD. The concentration of FAD-bound 64 NSMOB was determined to be 2 - 3 pM, as estimated by the absorbance drop at 450nm, which corresponds to FAD reduction in the stopped flow instrument at 15°C. Single-turnover kinetics were monitored by fluorescence excitation at 463 nm and emission at 520 nm. Lines passing through the data points depict the best exponential fits after logarithmic time averaging, non-time averaged data is presented in Appendix 6. The absorbance data in Figure 26 represents the NSMOB hydride-transfer reaction from B. E Qs UT>CM <3 5e 'm 1yj w 2© L i . Time/sec Time/sec Figure 27: Absorbance and Fluorescence Measurements of Hydride-Transfer Reaction. A) A340 absorbance. B) Fluorescence emission at 520nm. Both results depict the. A) NADH -> FAD hydride-transfer reaction kl = 56.7s-1 ± 3.7 B) FAD -> FAD hydride-transfer kinetics k2 = 8s"1 from the reaction of NSMOB with NADH and FAD. 65 NADH to oxidized FAD. The previous data determined the native SMOB hydride- transfer rate to be k = 49 ± 1.4 s_1 (Morrison, 2013). This research estimated the N- terminally tagged enzyme average rate constant to be k = 48 s'1. The data in Figure 27 were fit with a bi-exponential to give the hydride rate transfer constant to be kl = 56.7s'1 ± 3.7, AD-FAD transfer rate constant, k2 ~ 8s"1.In the reduction reaction of NSMOB in the presence of excess FAD, pyridine nucleotide to flavin hydride transfer occurs with a rate constant similar to that of the native enzyme, but this is followed by a slower NSMOB catalyzed flavin to flavin hydride transfer reaction of 8 s’1, which corresponds to the reaction of the FAD substrate in the double displacement mechanism (Figure 3A & B). 3.6 Efficiency of flavin-transfer from SMOB to SMOA 6 CMin Figure 28: Kinetics of eo 5 styrene epoxidation in the oc reaction catalyzed by g 4.5 NSMOB and NSMOA rate-limited by the FAD -> FAD hydride-transfer reaction. i 2.5 0 0.5 1 1.5 2 2.5 3 Time/sec 66 When NSMOA and styrene and oxygen are included a fluorescence increase corresponding to the accumulation of the C4a-hydroperoxyflavin intermediate at a rate limited by the flavin-flavin hydride transfer reaction at 8 s'1. Figure 28 shows the kinetics of NSMOB-FADoh, formation (k3 ~ 3s-l), rate-limited by the proceeding reduction and FAD-transfer reactions. In the absence of styrene NSMOA sequesters FAD as a stable C4a- hydroperoxyflavin intermediate as has been previously shown in the reaction with native SMOB in Figure 29 (Morrsion, 2013). This experiment showcases the hydride transfer (1), steady state reaction rate-limited by the kinetics of H202 elimination from NSMOA (2), and consumption of limiting reagent oxygen. 8.5 5 ------*------I------1------1------0 20 40 80 80 100 Time/sec 67 The preceding results have presented the binding affinity of oxidized FAD to NSMOB, which was determined by fluorescence-monitored equilibrium titrations to be -69 nM compared to native SMOB (Kd = 280 nM) at 4°C. The steady-state mechanism of NSMOB at 30°C was found to occur as a double-displacement reaction with a Vmax = 3.0 ± 0.4 pM-lmin-1, KmFAD = 6 ± 1 pM, and KmNADH = 4.8 ± 0.3 pM when compared to the sequential ordered mechanism of the native enzyme. Furthermore, the summary of the single-turnover results confirms that the N-terminal tag directly affects the rate limiting hydride-transfer reaction due to the increased affinity of flavin for NSMOB. The implications of these results will be discussed in the next section along with the proposed future research directions of engineered one- and two-component SMO systems. 68 DISCUSSION 4.1 N-terminally tagged SMO and protein engineering implications In light of new detailed biochemical and structural characterizations directed at uncovering the specific applications of naturally occurring and engineered two- component flavoenzymes, this research aims to expand on the utility of N-terminally tagged enzymes and N-terminally linked fusion proteins alike. The catabolism of toxic styrene in Pseudomonas putida (S12) is accomplished by the regio- and enantioselective oxidation of styrene using a two-component, styrene monooxygenase enzyme (SMO) in the presence of molecular oxygen, a quality that gives the SMO system immense biocatalytic potential (Huijbers, 2014). In an effort to enhance the efficacy of novel flavoprotein monooxygenases as biocatalysts, new recombinant DNA technology has allowed the improvement of substrate specificity due to key active site mutations, and alleviated the difficulty of protein expression and recovery through the use of N-terminal histidine tags (Kantz, 2005; Lin, 2012) In this research we investigated the effects of a 6- histidine N-terminal tag on the SMO reductase component, SMOB. We have distinguished the effects that this 20-amino acid moiety has on the catalytic FAD reduction, FAD binding affinity, and the hydride- transfer rate constant in light of what is currently known about the native enzyme and similar fusion proteins (Kant, 2005; Morrison, 2013; Tischler, 2010). 69 4.2 Isoalloxazine binding environment and SMOB refolding and stability In general the riboflavin-based coenzymes are bound to enzymes with high affinity and function in catalyzing the oxidations and reductions of aromatic compounds (Walsh, 2013), which enables a vast range of chemical transformations in many biosynthetic pathways. These particular flavoenzymes take part in oxidations involving amine and alcohol activating groups, with both the C(4a) and N(5) regions of the flavin coenzymes acting as sites for the covalent adduct formation that is characteristic of the two-component styrene monooxygenase enzymes (Walsh, 2013). We concluded that SMOB shows very little flavin specificity in catalytic turnover, however, both the yield of SMOB in the flavin-catalyzed protein-folding mechanism and the half-life of the folded protein are significantly dependent on the structure of the flavin catalyst. Studies on the stability of apo-SMOB generated using activated carbon concluded that the half-life of protein as significantly reduced and aided in its destabilization (Morrison, 2013). Although flavin is conclusively known to enhance the refolding of SMOB during purification and increase protein solubility, the type of AMP moiety present determines the overall increase in SMOB stability. These findings suggest a flavin-specific folding mechanism that favors the folding of SMOB with bound FAD over FMN or riboflavin. This is FAD-specificity in the folding mechanism of SMOB may be a type of regulatory mechanism that helps modulate the activity of SMOB in response to the concentration of FAD in the cell. 70 4.2.1 Flavin-binding affinity and SMO catalytic mechanisms The observed changes in FAD- binding affinity directly impact the steady-state kinetic mechanism, which changes from an ordered sequential to double displacement in the case of NSMOB. Native SMOB binds flavin with a Kd of 356 ± 30 nM at 4°C, with that affinity decreasing significantly at higher temperatures (Morrison, 2013). NAD* fmrnrnrnmm FAD,V«d \ Jg> F m m . NAD* NADH ^ FAD, NAD* m m m m F A D * >FAEW Jmmm ^ FAD,* km m fm m n n n ■ A a •— NADH mmmUADH FAD,. g— JLFWrT" J— t-TT* t f FA0- / f FAD„» f Figure 30: NSMOB Interchange of Sequential and Double Displacement Reaction Mechanisms based on Kd. The sequential mechanism (left) and double displacement mechanism (right) are related through the equilibrium dissociation constant that describes the reversible binding of FAD to apo- NSMOB. The Km apparent equations used in the steady-state determination of the SMOB catalytic mechanism are defined below, and the major difference is denoted in blue. 71 Our hypothesis that the N-terminal tag confers SMOB with a differential flavin binding environment was investigated using steady-state kinetic experiments and non linear least squares curve fitting to provide estimates of the Km of NADH, Km of FAD and Vmax ofN-SMOB at 3.0 |aM, 6.0 pM, and 4.8 pMs-1, respectively at 30°C (Table 2). Figure 30 represents a schematic interpretation of the relationship between Kd and the change in catalytic mechanism when flavin binding affinity is increased due to active- or non-active site changes. In detail, the above figure illustrates the flavin-flavin reaction that is shown to be the predominant mechanism of flavin reduction for the N-terminally tagged version. Based on this research we can assume that other genetically engineered or naturally-fusion proteins that N-terminally link both the flavin reduction and styrene epoxidation reactions on the same polypeptide exhibit similar changes in catalytic reactivity, which may be attributed to the increased flavin binding affinity (Tischler, 2013). Recent crystallographic data confirmed that the wild-type SMOB enzyme is a homodimeric peptide with structural and mechanistic similarities to the well characterized, PheA2, a flavin reductase enzyme from Bacillus thermoglucosidasius A7 (Morrison, 2013; Heuvel, 2004). The PheA2 reductase binds FAD with a Kd of 9.8 nM, while the HpaC unit binds FAD in the uM range (Kim, 2007). In nature, both the reductase and monooxygenase subunits interact as dimers to catalyze the epoxidation of styrene to (S) - styrene oxide (Heuvel, 2004). Interestingly, PheA2 also catalyzes the 72 reduction of FAD using a BiBi sequential mechanism during low concentrations of the catalytic flavin (Heuvel, 2004). Despite the similarities in active site composition and the presence of a pHBH fold, we now known that the FAD cofactor shows differential binding between PheA2 and HpaC structures, which would be amplified in the presence of an engineered N-terminal tag as is the case for NSMOB. We expect that new fusion enzymes and N-terminally tagged enzymes, like NSMOB, will also display key similarities due the importance of the N-terminus as a regulatory domain of FAD reduction and subsequent transfer to the SMOA epoxidase (Morrison, 2013). Furthermore, the recent discovery of a unique styrene monooxygenase (StyA2B) from Rhodococcus opacus 1CP represents a new group of class E- flavoproteins that are naturally found with the reductase and epoxidase components on a single polypeptide (Tischler, 2009). This data in conjunction with our results provides information that will further elucidate the catalytic mechanisms of naturally occurring and engineered one- and two- component enzymes as valuable biocatalysts. The comprehensive advantages and disadvantages of each system with regards to the stability and catalytic activity of N- terminally linked fusion proteins and their respective substrates should be evaluated in future research. 4.3 SMO Rate-limiting hydride-transfer step We found that the hydride-transfer reaction from NADH to FAD in the active site NSMOB at 56.7s-1 ± 3.7 is relatively unchanged when compared with the native enzyme, which possesses a kl= 49 ± 1.4 s_l (Morrison, 2013). Due to the significant change in 73 catalytic mechanism and increased FAD-binding affinity we expected that pre-steady state kinetics would also be affected by the addition of an N-terminal tag. However, this data effectively resolves the rate limiting step for NSMOB to be the flavin-flavin electron-transfer step following the hydride transfer, which is significantly different than the wild-type enzyme. Figure 31 illustrates how the styrene epoxidation reaction of SMOA is rate-limited by the pyridine nucleotide - flavin hydride transfer in native SMOB (Morrison, 2013) k, ~ 50s*1 k, ~ 100s'1 Styrene Styrene ,, „ _ NADH MAD* «— ; ' 0' it4e — b HjO p i ■ V / ■ V ./ ■ V / SMOB SMOB SMOB | Oj * SMOA | i ■■■ It. SMOA-FAD^ SMOA*, Figure 31: Native SMOB catalytic mechanism including corresponding rate constants, substrates and catalysis. Putative enzyme binding sites are denoted in blue. Figure 31 summarizes the important steps in the reduction of FAD by native SMOB showing that the rate limiting-step for this enzyme is the kl = 50 s'1. The oxidation rate of reduced flavin is dependent on the concentrations of oxygen and oxidized flavin present during the experiment, and our studies utilized parameters that were previously defined for this type of reaction (Kantz, 2005). In light of the new studies confirming the presence of an SMOB-SMOA complex we can assume that the presence of oxygen and oxidized flavin may reduce SMOBs capacity to effectively 74 reduce and transfer flavin to SMOA. In the case of the native reductase a transient complex to facilitate the translocation of reduced flavin from SMOB active site to apo- SMOA has been elucidated (Morrison, 2013). However, in the presence of an N-terminal tag, NSMOB may have to rely on the diffusion of reduced flavin and slower flavin-flavin hydride transfers to efficiently complete its reductive half-reaction shown in Figure 32. k1Pj ~ 50s'1 ~ 8s"1 k3N ~ 3 s'1 ™ 1.5 s 1 Figure 32: N-terminally tagged SMOB catalytic mechanism including corresponding rate constants (kN), substrates and catalysis. Putative enzyme binding sites are denoted in blue. Figure 32 summarizes the kinetics of the dislocation of reduced flavin from NSMOB (k4 ~ 3 s'1) to be much slower than that of the native enzyme due to the increased flavin binding affinity, which also affects the NSMOA-FADoh, formation (k3 ~ 3 s-1), rate-limited by the proceeding reduction and FAD-transfer reactions. In 75 comparison both versions of the reductase, when there is excess oxidized flavin present, still perform a fast initial reduction that appears to be unchanged despite the presence of an N-terminal tag. Additionally, this fast pyrimidine nucleotide to oxidized flavin electron transfer (Icni = 50 s'1) is followed by the steady-state turnover of oxidized flavin after the epoxidation of styrene to form the SMOA-FADOH intermediate, which can then be stabilized by apo-SMOB thus starting the cycle over again and preventing the production of reactive oxygen species. In conclusion, the modulatory impact of the N- terminus on the reductase enzyme is greatly affected by the addition of a new peptide as it causes a complete change in the catalytic mechanism and FAD binding affinity. Based on these observations and new data we should expect to see similar changes in other N- terminally linked flavin monooxygenase systems, regardless if they are engineered or naturally occurring. 4.4 SMO Flavin-transfer mechanisms andprotein-protein interactions While no evidence of an SMOB-SMOA complex has been visualized in present structural studies from 4-hydroxyphenylacetate-3-monooxygenase or SMO (Morrison, 2013), experiments indicate that the coupling efficiency of the reductive and oxidative half-reactions is higher than what should be expected of a purely diffusive reaction (Morrison, 2013). Additionally, the Vi life of SMOB in the absence of its bound flavin during the FAD-transfer reaction is ~21 fold less than its V2 life in the presence of FAD, supporting a protein-protein interaction between apo-SMOB and the SMOA peroxide intermediate (Morrison, 2013). The stabilizing effect of FAD has been continuously 76 recognized in the expression and purification of high yields of soluble, active SMOB reductase (Kantz, 2005), and our studies have implicated high amounts in excess FAD in the successful purification of NSMOB also. Previous studies determined that in the absence of oxidized flavin and in an excess of NSMOA, the epoxidase binds the majority of flavin as a stable C4a-hydroperoxy intermediate that facilitates the catalytic conversion of styrene to styrene oxide (Morrison, 2013; Bach, 2014). In our pre-steady state analysis of NSMOB we determined that while excess flavin does allow the visualization of the highly fluorescent intermediate, it also prevents our accurate determination of the rate- limiting step in the case of the N-terminally tagged enzyme. The increased flavin binding affinity directly regulates the hydride-transfer reaction and causes the prevalence of a much slower FAD-FAD electron transfer step and subsequent reduced flavin translocation to the active site of apo-SMOA. This slow step may be imperative for NSMO catalysis because the 20-amino acid tag could functionally inhibit the direct interfacing of each subunit for efficient flavin transfer. Moreover, the histidine tagged protein has a significantly increased FAD-binding affinity compared with the native enzyme is consistent with the observed change in steady-state mechanism from sequential to double displacement, which more closely relates NSMOB to the phenol hydroxylase reductase, PheA2 from Bacillus thermoglucosidasius A7 and styrene monooxygenase fusion proteins, which similarly follow a ping-pong mechanism (Heuvel, 2004; Kim, 2007; Tischler, 2013). The styrene epoxidation and reaction of wild-type SMO is rate-limited by the kinetics of the pyridine 77 nucleotide to FAD hydride transfer reaction, and contrasts the rate of styrene epoxidation, which is rate-limited by the flavin-flavin hydride transfer in NSMOB. This work highlights the significance of the N-terminus of the reductase in defining the FAD- binding affinity and kinetics of FAD-transfer in two-component and naturally-occurring fusion proteins and will be an important consideration in the design of engineered SMO fusion proteins for biocatalysis. 4.5 Engineered Flavin Monooxygenases as Efficient Biocatalysts Previous studies focused on the intrinsic catalytic activities of microorganisms that are able to detoxify their environment and transform styrene into biomass (Reetz, 2013; Ceccoli, 2014). It is the expanded understanding of the biochemical mechanisms by which styrene is degraded that will prove essential to determining its impact on human on health and viability of styrene monooxygenase and other similar systems as effective biocatalysts. Our work on the unique components of the styrene monooxygenase system of Pseudomonas putida S12 highlights the essential regulatory nature of the N-terminus of engineered one- and two-component flavoenzymes in toxin metabolism. The addition of the 6-histidine N-terminal tag did not result in a modification of the rate limiting step, allowing the reaction to remain coupled, which in turn circumvents the production hydrogen peroxide and superoxide species in the presence of SMOA, a characteristic that is highly desirable when engineering safe biocatalysts. 78 Despite the decreased catalytic turnover of reduced FAD, the increased binding affinity and change to a double displacement mechanism would prove valuable in situations where decreased amounts of free catalytic flavin are present (Lin, 2012). The addition of the N-terminal tag greatly reduced the manpower and time needed to purify large amounts of stable and active SMOB enzyme. 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[BSA] (pg/mL) Appendix 3: BSA assay standard curve using bovine serum albumin as a reference. Figure represents the best BSA curve fit for NSMOB to determine total protein concentration. 87 Total NSMOB (mg) 22 Total Activity (jimol.mln1) 1320 Specific Activity (p.mol.mg1m in1) 58.8 Appendix 4: NSMOB BSA assay standard curve using bovine serum albumin as a reference. Figure represents the best BSA curve fit for NSMOB to determine total protein concentration. NSMOB Steady State Data 12 10 8 13 4 2 0 0 10 20 30 40 50 [FAD] uM Appendix 5: Steady State Kinetic Plots of NSMOB with NADH and FAD. The reaction of NSMOB with increasing concentrations of NAD and FAD is shown. Data traces correspond to reactions run at 2|iM, 3|uM, 5|jM, 7|liM, 15|liM, 25|iM, 35|iM and 50|uM. Each plotted point corresponds to an average value of 3 independent runs with one standard deviation denoted by the Y-error bars. Lines passing through the data points correspond to the Michaelis-Menten fit using KaleidaGraph® software, estimates of the Km of NADH, Km of FAD and Vmax of N-SMOB parameters were derived from this plot. 89 0.88 0.86 0.84 0.82 0.078 0.78 0.1 0.2 0.3 0.4 0.5 0.6 0.7 Time/sec Appendix 6. Single-turnover NSMOB stopped-flow data. Single-turnover, stopped- flow experiments examined the pre-steady state kinetics ofN-SMOB. (Blue) represents the oxidation of NADH at 340nm, (Green) fluorescence excitation at 450nm and emission at 520nm, and (Red) reduction of FAD at 450nm.