biochemical characterization of n-terminally tagged
TRANSCRIPT
BIOCHEMICAL CHARACTERIZATION OF N-TERMINALLY TAGGED STYRENE MONOOXYGENASE FROM PSEUDOMONAS
A thesis submitted to the faculty of San Francisco State University
In partial fulfillment of The Requirements for
The Degree
Master of Science In
Chemistry: Biochemistry
by
Nonye Nwa-Niaf Okonkwo
San Francisco, California
May 2015
A S
• O U
Copyright by Nonye N. Okonkwo
2015
CERTIFICATION OF APPROVAL
I certify that I have read The Biochemical Characterization o f N-Terminally Histidine
Tagged Styrene M onooxygenase from Pseudomonas by Nonye N w a-N iaf Okonkwo, and that
in my opinion this work meets the criteria for approving a thesis submitted in partial
fulfillment o f the requirements for the degree: Master o f Science in Chemistry: Biochemistry
at San Francisco State University.
Professor o f Biochemistry
Weiming Wu, PhD Professor o f 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..........................11.2 Microbiology of styrene degradation.............................................................51.3 Styrene catabolic and detoxification pathway..............................................111.4 One- and two-component styrene monooxygenases.................................... 141.5 Catalytic mechanisms of styrene monooxygenases......................................181.6 Styrene monooxygenase structure.................................................................251.7 Naturally-occurring and artificially engineered fusion proteins................ 271.8 Putative complexes..........................................................................................301.9 Transition to thesis research............................................................................32
Methods............................................................................................................................... 34
2.1 Protein expression and purification...............................................................342.2 Activity and purity assays..............................................................................362.3 Time-dependent activity studies.................................................................... 392.4 Steady-state kinetic analysis ofNSMOB..................................................... 402.5 Estimation of equilibrium dissociation constants........................................ 432.6 Pre-steady-state kinetics.................................................................................44
Results................................................................................................................................. 46
3.1 Flavin-refolded SMOB experiments with FAD, FMN, and riboflavin....463.2 Flavin-refolded SMOB time dependence studies........................................ 523.3 NSMOB mechanistic studies..........................................................................563.4 Estimation of equilibrium dissociation constants by fluorescence Monitored Titrations..............................................................................................60
vi
3.5 NSMOB single-turnover studies................................................................... 613.6 Efficiency of oxidative- and reductive half-reactions.................................65
Discussion........................................................................ 68
4.1 N-terminal tags and implications in protein engineering............................ 684.2 Isoalloxazine ring environment: SMOB refolding and stability................ 694.3 SMO Rate-limiting hydride-transfer s tep ....................................................724.4 SMO Flavin-transfer mechanisms and protein-protein interactions..........754.5 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 SM O B..................................... 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 o f 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).
Biologically styrene acts as a human cellular
membrane disrupter and manifests its intracellular
effects through cytochrome p450 isofroms, which
convert styrene to SO and 4-vinylphenol, which
have biological activity as carcinogens and
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
oVI
Styrene Styrene-7, 8- Oxide
Figure 1: Chemical Structure of Styrene and (Sj-Styrene-7, 8- Oxide.
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 o f 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 o f 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«
G lticuro n icta im d su tfa to c o 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 o f 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
styrenestyrene
monooxygenase
styCstyrene oxide
styrene oxide phenylacetaidehyde
styDphenylacetaidehyde
dehydrogenase
OH ^ ^ ^SCoATCA Cycle
O phenylacety! coAttgase
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 o f 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
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
styrene detoxification pathway in
Pseudomonas sp. converts styrene to (S)-
styrene-oxide by using an NADH-dependent
reductase (SMOB) and an FAD-specific
monooxygenase (SMOA) (Van Berkel, 2006; Montersino, 2011) as seen in Figure 4. The
12
Figure 4: Styrene Monooxygenase (SMO) Catalysis. SMOB (pink) catalyzes NADH-specific reduction of FAD and SMOA (green) performs subsequent FAD-dependentepoxidation of styrene to form (S)- styrene oxide (Hartmans, 1990).
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
' - * 1.4 One- and two-componentid J\ / monooxygenase systems
R'
R ' ohH ! ° HI . OH
X X ^ v ™ M Flavin-dependentH O i f R R iR
/ HO ^ vnJ \{4) enzymes are highly abundant in
/ \
1 XR R’________________ x-s,s,se.n,ft____________ as indispensable tools in the
nature and are widely regarded
Figure 5: C4a-hydro-peroxyflavin Intermediate Acts as a Precursor for Many Important Reactions Catalyzed by Flavin-Dependent Monooxygenases. (1) Epoxidation of C=C double bonds, (2) Baeyer- Villiger oxidation of ketones, (3) hydroxylation of phenols, (4) heteroatom oxygenation, (5) aldehyde oxidation, (6) halogenation reactions (Holtmann, 2014).
synthesis of biologically active
compounds (Chaiyen, 2012).
The remarkable enantio
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 o f styrene monooxygenases
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 halfreaction (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
A
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 o f 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 o f 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
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
20
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 o f 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 o f 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
f cMo rt. - "vSv^ version of SMOB was^ e \ \ ' » I J r
H ' v - \V iA jX used for x-ray
I * I 1 x ■ '% ' i t i iYj \ i '> «vf crystallography, the
crystal structure of the
20kDA SMOB unit
was solved with a 2.2
A resolution. It was
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
Figure 9: Overall Structure of N-terminally Tagged Styrene Monooxygenase A. (A) NSMOA monomer. Domain A is colored blue and domain B green. NSMOA secondary structure. (B) NSMOA dimer colored as in panel A. (Ukaegbu, 2010).
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 StyA2Bsiymrt* 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 o f
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 S I2 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 o f soluble SMOB from inclusion bodies andflavin dialysis
120 ml o f 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
H* II O 1 — e rate equation for timedependence 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 o f 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 o f 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 o f 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
,. [N A D H ] [FAD]r,W l/°»' _____“ ~ K m™ + (NADH] “ K T + [**£>]
Kmapp Equations
k T wadh] K ^ \ fad]K T h + \n a d h ] A m k ™ + [ f a d )
v v% [ f a d \K°pp + [FAD]
Equation 2: KineticFitting Equations. V max
apparent and Km apparent equations. The apparent Km and V m a x parameters were used in GraphPad Prism software global- fitting analysis.
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 o f 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 - J (K “ pr>+{FAD]tow! + jS M O B ]^ y -4[FAD]totai[SMOA]R (£2 -e i)
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
/t&m/ - r # ’
wild-type SMOB produces a
high yield of the reductase in the
form of insoluble inclusion
bodies, which can be denatured
in urea and refolded using
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
Figure 12: Flavin-mediated refolding of SMOB in 8M urea.
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.
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
Figure 13: FlavinStructure. Riboflavin, Flavin Mononucleotide (FMN),and Flavin Adenine Dinucleotide (FAD).
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
100Ic SO1CL 60
I 40
20
0FAD FMN Riboflavin
Flavin
Flavin-RefoldedSMOB
Total Protein [mg)
Total Units of Activi ty (pmole/rain)
Specific Activity (mU/mg)
FAD 124 + 25 1.15 9.2FMN 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 o f SMOB refolded in the presence o f FAD, FMN, and riboflavin
I Li I_______ I_________ __J0 2 4 6 8 10
Time at 4°C (Days)
Figure 18: Half-Life of SMOB in the Presence of FAD, FMN, and Riboflavin at 4°C. FAD (red), FMN (green), and riboflavin (blue).
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 o f SMOB
A. B.
Wavelength <nm) in m)
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 o f 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 o f 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: RibbonStructure of N-terminally Tagged FAD-boundSMOB interacting with a Nickel Affinity Column agarose bead.
OWe 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 o f 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) pM
Km (FAD) pM
VmaxpMmin"1
Ks (NADH) pM
Sequential 3.0 ±0.7 6.0 ± 1.6 4.8 ±0.4 1.0'7± 1.3
DoubleDispl.
3.0 ±0.5 6.0 ± 1.0 4.8 ±0.3 N/A
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
c£
* 2 - 3* 5i 7 ’ 10 *• 12 J 20
Figure 23: Global fit NSMOB Double Displacement Mechanisms. The graphs displayvelocities from reactions in which both NADH and FAD (x-axis)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 D isplacem entNull 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 o f 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
| 450000008 400000 c1 350000£US
e| 250000o3£
200000
150000 500 1000 1500 2000 2500 3000 3500 (FAD) total <nM)
Fig 24: Fluorescence titration monitoring of FAD to NSMOB. The fit equation provided an accurate estimate of the equilibrium dissociation constant of NSMOB to be ~ 69 nM at 4°C.
61
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 Equation6: Steady-state NSMOB kinetic data taken on a stopped-flow device. A340 exponential and line equation fit.
0.7950.08 0.16 0.24 0.32 0.4 0.48 0.56 0.64
Time/sec
4 3 4 0 (f) = 4340 exp (~kt} v t + 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. Singleturnover, 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.
EsQCMUT><3e5'm1yj
w©2L 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 o f 8 s’1, which corresponds to the reaction of the FAD substrate in the double
displacement mechanism (Figure 3A & B).
3.6 Efficiency o f flavin-transfer from SMOB to SMOA
6
g 4.5
CMin
coeo 5
i
NSMOB and NSMOA rate-limited by the FAD -> FAD hydride-transfer reaction.
Figure 28: Kinetics ofstyrene epoxidation in the reaction catalyzed by
2.50 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 o f 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 o f 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).
FAD,V«dNAD* fmrnrnrnmm
\ Jg> F m m . NAD* NADH ^ FAD,
NAD* mmmm F A D *
>FAEW Jmmm ^ FAD,* km m fm mn n n
■ A a•— NADH mmmUADH FAD,. g—
J L FWr T " J— t - T T * tf 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 k l= 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
| O j * SMOA |i ■ ■ ■ I t . SM OA-FAD^ S M O A *,
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 o f 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 o f 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. In addition to providing insight into
the structure and mechanism of SMO in the styrene catabolic and detoxification pathway,
this work and surrounding research efforts provide the foundation needed to engineer
new functions and substrate specificity in flavoproteins as they are developed as
biocatalysts for the enantioselective synthesis of fine chemicals, pharmaceuticals, and
other bioactive compounds (Reetz, 2013).
78
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APPENDIX
oo
Fraction #
Appendix 1: NSMOB FPLC Fraction Plot. (Red) conductivity, (blue) gradient pump, (green) UV are plotted against fraction number. 83.4 protein fractions were collected and fractions 55-75 were pooled based on UV absorbance. NSMOB was eluted using a linear gradient of imidazole increasing from 10 to 250 mM.
Appendix 2: NSMOB SDS-PAGE Analysis. Lane 1: post sonication supernatant; Lane 2: post sonication pellet; Lane 3: FPLC fraction 14 ‘flow through’; Lane 4: Fraction 57; Lane 5: Molecular weight standards; Lane 6: Fraction 60; Lane 7: Fraction 68; Lane 8: Pooled fractions 55-7; Lane 9: N-SMOB glycerol stock.
[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
10
8
12
13
4
2
00 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
0.078
0.88
0.86
0.84
0.82
0.780.1 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.