Download Human Cytochrome P450 2E1 - KU ScholarWorks

Human Cytochrome P450 2E1:
Functional Comparison to Cytochromes P450 2A13 and 2A6
by
Melanie A. Blevins
B.S., Graceland University, 2002
Submitted to the Department of Medicinal Chemistry and the Faculty of the
Graduate School of the University of Kansas in partial fulfillment of the
requirements for the degree of Master’s of Science.
Dissertation Committee:
Major Professor
Committee
Date Defended: April 24, 2008
The Thesis Committee for Melanie Blevins certifies
that this is the approved Version of the following thesis:
Human Cytochrome P450 2E1:
Functional Comparison to Cytochromes P450 2A13 and 2A6
Committee:
_______________________________
Chairperson*
_______________________________
_______________________________
ii
Abstract
The cytochrome P450 (CYP) superfamily of enzymes plays the
predominant role in human phase I xenobiotic metabolism. These enzymes
participate in the metabolism of a greater part of the drugs in present clinical
use and have been linked to the activation of carcinogens and other toxins.
The CYP2 family, in particular, is known for it extensive Phase I metabolism
of a majority of the xenobiotic compounds [1]. The goal of this project is to
determine the structural foundation for the substrate selectivities of the
CYP2A6 and CYP2A13 enzymes versus CYP2E1. Because these enzymes
metabolize both common, as well as unique, small molecule substrates, it is
likely that only few key residue-substrate interactions are responsible for
those metabolic capabilities that differ between them.
Amino acid residues in regions of the CYP2E1 protein likely to contact
ligands and that differ between CYP2E1 and the CYP2A enzymes were
examined by site-directed mutagenesis.
The resulting mutated CYP2E1
proteins were characterized for their ability to hydroxylate the reportedly
selective CYP2E1 substrates p-nitrophenol (pNP) [2] and chlorzoxazone
(CZN) [3], but none showed significant differences in activity from the
CYP2E1 wild type enzyme.
iii
However, in contrast to previous literature reports [4], both CYP2A6 and
CYP2A13 were observed to metabolize both CYP2E1 substrates pNP and CZN
with catalytic efficiencies equal to or greater than CYP2E1 (Table 1). These
unexpected activities of the CYP2A enzymes with CYP2E1 substrates
demonstrate that the human CYP2A and CYP2E enzymes are more
functionally similar than previously believed.
Table 1: pNP and CZN kinetic parameters for CYP2E1, CYP2A13, and CYP2A6.
p-Nitrophenol
Chlorzoxazone
CYP450
kcat
(min-1)
Km
(µM)
kcat/Km
(µM-1min-1)
kcat
(min-1)
Km
(µM)
kcat/Km
(µM-1min-1)
2E1
15.9 ± 0.4
75.8 ± 4.4
0.21
5.1 ± 0.4
105.5 ± 22.9
0.05
2A13
30.3 ± 1.3
62.7 ± 7.4
0.48
22.7 ± 1.1
64.8 ± 10.4
0.23
2A6
52.6 ± 2.4
135.8 ± 13.7
0.39
4.7 ± 0.3
100.7 ± 15.1
0.07
References
1
Rendic, S. and Di Carlo, F. J. (1997) Human cytochrome P450 enzymes: a
status report summarizing their reactions, substrates, inducers, and inhibitors.
Drug Metab Rev 29, 413-580
2
Koop, D. R., Laethem, C. L. and Tierney, D. J. (1989) The utility of p-nitrophenol
hydroxylation in P450IIE1 analysis. Drug Metab Rev 20, 541-551
3
Peter, R., Bocker, R., Beaune, P. H., Iwasaki, M., Guengerich, F. P. and Yang, C. S. (1990)
Hydroxylation of chlorzoxazone as a specific probe for human liver cytochrome P-450IIE1.
Chem Res Toxicol 3, 566-573
4
Zerilli, A., Ratanasavanh, D., Lucas, D., Goasduff, T., Dreano, Y., Menard, C., Picart,
D. and Berthou, F. (1997) Both cytochromes P450 2E1 and 3A are involved in the Ohydroxylation of p-nitrophenol, a catalytic activity known to be specific for P450 2E1.
Chem Res Toxicol 10, 1205-1212
iv
Acknowledgements
I would like to thank all of the people who helped me along the way
during my graduate career. I would like to acknowledge all of the guidance
from Brian Smith and Lena Zaitseva in helping with the optimization of the
CYP2E1 purification protocol. I would also like to thank Patrick Porubsky for
the purification of NADPH-P450 oxidoreductase, CYP2A13, and CYP2A6. I
would like to thank all of the past and present members of the Scott lab,
Natash DeVore, Naseem Nikaeen, Eric Carrillo, Anu Metha, Agnes Walsh,
Kathleen Meneely, and Linda Blake for their support and encouragement over
the last few years.
Thank you to my committee members Dr. David Benson and Dr. Sunil
David for taking the time to review my research. A humongous thank you to
my advisor, Dr. Emily Scott, for her continous patience, and guidance through
out my graduate career. A special thank you to the NIH RR17708 and
GM076343 grants for funding.
Finally, I’d like to offer a huge thank you to my family and friends who
have counseled me through my graduate career. You all have contributed
greatly to my success here at the University of Kansas. Without all of your
support I would have never made it this far.
v
Table of Contents
Page
Abstract
iii
Acknowledgements
v
List of Figures
viii
List of Tables
x
List of Schemes
xi
Chapter 1. Introduction to Cytochromes P450
1
Introduction
1
Xenobiotic Metabolism
2
Cytochromes P450
4
CYP450 Enzymes: Organization of Isozymes and Nomenclature
8
CYP450 Structure and Active Site Topography
10
The Catalytic Cycle
13
CYP450 2 Family: Substrate Overlap and Diversity
16
CYP2A6
19
CYP2A13
21
CYP2E1
23
Enzyme Kinetics
25
References
28
Chapter 2. Project Goals, Hypothesis, and Design
37
Project Goals and Hypothesis
37
Project Design
38
References
42
Chapter 3. Site-Directed Mutagenesis, Expression, and Protein
Purification
Introduction
44
44
vi
Methods
56
Site-directed mutagenesis
56
Purification of Plasmid DNA
59
Restriction Enzyme Digestion
60
Protein Expression
62
Protein Purification
63
Results
64
Conclusions
71
References
72
Chapter 4. Characterization of 2E and 2A Proteins Using
Chlorzoxazone and p-Nitrophenol Hydroxylation Assays
73
Introduction
73
Methods
80
p-Nitrophenol Hydroxylation Assay
80
Chlorzoxazone Hydroxylation Assay
82
Results
83
p-Nitrophenol Metabolism Assay
84
Chlorzoxazone Metabolism Assay
90
Conclusions
94
References
97
Chapter 5. Conclusions
100
References
111
vii
List of Figures
Figures
Page
1-1.
Carbon monoxide difference spectrum
7
1-2.
CYP450 enzyme heme moiety
7
1-3.
Crystal structure of CYP2A13: substrate recognition sites
11
1-4.
CYP450 enzymes involved in phase I metabolism
17
1-5.
CYP2A6 substrates
20
1-6.
CYP2A13 substrates
22
1-7.
CYP2E1 substrates
24
1-8.
Michaelis-Menten kinetics
27
2-1.
Amino acid sequence alignment of CYP2E1, CYP2A13, and
CYP2A6
40
3-1.
Site-directed mutatgenesis strategy
46
3-2.
QuikChange II site-directed mutagenesis strategy
49
3-3.
Schematic of Ni-NTA metal affinity column chromatography
53
3-4.
Schematic of ion-exchange column chromatography
55
4-1.
The hydroxylation of p-nitrophenol
76
4-2.
The hydroxylation of chlorzoxazone
78
4-3.
Comparison of CYP450 p-nitrophenol activities
85
4-4.
Comparison of Michaelis-Menten kinetics determined by both
visible colorimetric and HPLC UV detection methods
4-5.
87
Overlay of enzyme kinetics for p-nitrophenol metabolism by
CYP2E1, CYP2A6, and CYP2A13.
89
4-6.
Comparison of CYP450 chlorzoxazone activities
91
4-7.
Overlay of enzyme kinetics for chlorzoxazone metabolism by
CYP2E1, CYP2A6, and CYP2A13
93
viii
5-1.
Overlay of crystal structures of CYP2E1 and CYP2A13
104
5-2.
Crystal structure of CYP2E1: substrate recognition site 1
106
5-3.
Crystal structure of CYP2E1: substrate recognition site 2
109
List of Tables
Tables
1-1.
Page
Human CYP2 family enzymes and their known locations,
reactions, and inducers
18
3-1.
Thermal cycling parameters used for mutagenesis reactions
58
3-2.
Restriction enzyme reaction conditions
61
3-3.
The physical characteristics of designed oligonucleotides
65
3-4.
Site-directed mutagenesis results
66
3-5.
Characterization of purified CYP2E1 proteins by UV/Vis
spectroscopy and CO difference spectra.
4-1.
Comparison of p-nitrophenol activity determined by visible
colorimetric and HPLC UV detection methods
4-2.
93
Comparison between CYP2E1 mutant protein activities for
p-nitrophenol and chlorzoxazone
5-1.
89
Chlorzoxazone kinetic parameters determined for CYP2E1,
2A6, and 2A13
4-4.
87
p-Nitrophenol kinetic parameters determined for CYP2E1,
2A6, and 2A13
4-3.
69
96
p-Ntirophenol and chlorzoxazone kinetic parameters for
CYP2E1, 2A13, and 2A6.
101
ix
List of Schemes
Schemes
1.1.
The catalytic cycle of cytochromes P450
Page
14
x
Chapter 1
Introduction to Cytochrome P450 Enzymes
Introduction
Metabolism can be described as the chemical and physical processes
occurring within a living organism, involved in the maintenance of life [1]. In
humans, a large portion of metabolic energy is involved in energy production
(catabolism) and protein and nucleic acid biosynthesis (anabolism).
Nonetheless, xenobiotic metabolism plays a crucial role in preserving
homeostasis. Because humans are continually exposed to a variety of foreign
compounds, the metabolic conversion and subsequent removal of these often
lipophilic and toxic compounds from the body is an important process.
Over the years, the human body has evolved the ability to defend itself
against lipophilic environmental toxins that otherwise persist in cells. The
primary defense is the use of enzymes that metabolize these lipophilic foreign
compounds into more polar molecules that can easily be excreted. Enzymes
catalyze the majority of these chemical transformations in the liver and many
other extrahepatic tissues, including the kidneys, brain, respiratory tract, and
GI tract [2].
1
Consideration of the metabolism of a drug is an integral part of
developing a drug for clinical use. It is one of the four main pharmacological
considerations
when
administering
a
drug:
absorption,
distribution,
metabolism, and excretion [3]. If a drug is cleared too slowly or too quickly,
the drug will not be maintained within the therapeutic window, causing
treatment to fail due to either lack of therapeutic effect or associated toxicity
[4]. It is also important that the drug be metabolically converted into
metabolites that are relatively nontoxic. Accordingly, prior to drug approval,
knowledge about the metabolic pathways and disposition of the drug is
required.
Xenobiotic Metabolism
Xenobiotic compounds are metabolized by a variety of enzymes that
first modify the parent compound to a more water-soluble metabolite and then
tag the metabolite for subsequent export from the cell. If the lipophilic parent
xenobiotic is not metabolized into a water-soluble metabolite that can be
readily excreted, it will linger in the body generating a sustained biological
response [5]. In addition, the metabolism to water-soluble metabolites not only
increases the rate of excretion, but also typically detoxifies and inactivates
2
compounds. As a result, drug metabolism is generally regarded as a
detoxification process [6].
Nonetheless, it is not accurate to presume that the metabolism of all
xenobiotics is always inactivating. Some compounds are metabolized into
biologically active metabolites. Occasionally, these metabolites themselves
are instrumental in eliciting the toxicological or pharmacological effect(s) [2].
When a parent compound is inactive and must be metabolically converted to
the active metabolite [7], the parent compound is termed a prodrug.
Sometimes the metabolite is not only active, but also responsible for harmful
consequences (e.g., carcinogenicity, tissue necrosis, teratogenicity).
Xenobiotic metabolism is separated into two categories: phase I and
phase II reactions [6, 8]. The defining feature of phase I (functionalization)
reactions is the introduction or unmasking of a functional group(s) (e.g.
COOH, OH, SH, NH2) to increase the water solubility of a lipophilic molecule
[2].
Phase
I
reactions
include
oxidative,
reductive,
and
hydrolytic
transformations. These reactions do not always introduce sufficiently
hydrophilic functional groups to bring about elimination or completely alter the
pharmacologic properties of the drug. They do, however, typically create a
functional group “handle” that can facilitate a subsequent phase II reaction(s).
3
The defining feature of phase II reactions is the conjugation of small,
hydrophilic, and ionizable compounds (e.g., glutathione, sulfate, glycine, and
other amino acids) to the xenobiotic compound [9]. These small compounds
are attached to the introduced or inherent functional groups of phase I
metabolites to generate a more polar and more readily excreted conjugated
product. These conjugated metabolites are generally absent of biological
activity and toxicity [5]. Therefore, both phase I and phase II reactions play
important roles in determining the pharmacokinetics of drugs and the activities
and persistence of xenobiotics.
Cytochromes P450
Cytochromes P450 are a superfamily of heme proteins that are the
primary phase I metabolizing enzymes, responsible for the metabolism of
both endogenous and exogenous compounds [10]. Cytochrome P450
enzymes (CYP450s) are found in all kingdoms of life and exhibit extraordinary
diversity in their reaction chemistry (e.g., aromatic and aliphatic hydroxylation,
epoxidation, N-, O-, and S-dealkylation, oxidative deamination, N- and Soxidation, dehalogenation). The substrates and reactions of CYP450s are
extremely diverse, including drug metabolism, the biosynthesis and
4
metabolism of fatty acids, steroids, and vitamins, detoxicification of
carcinogens and pesticides, and the activation of procarcinogens [11].
There are 57 CYP450s in the human genome, 42 of which have known
catalytic
activities.
Fifteen
CYP450s
are
associated
with xenobiotic
metabolism, fourteen are crucial in steroidogenesis, five catalyze the
metabolism of eicosanoids, four metabolize vitamins (A and D), and four have
fatty acids as their substrates [12]. A few of the xenobiotic-metabolizing
CYP450s can also catalyze the oxidation of steroids and fatty acids but these
functions do not appear to be critical to homeostasis (e.g., lauric acid 11hydroxylation by CYP2E1 [13] ).
Although soluble forms exists in bacteria and other lower organisms, in
mammals cytochromes P450 are primarily membrane-associated enzymes,
located in either the inner membrane of the mitochondrion or the smooth
endoplasmic reticulum [14]. These enzymes are highly expressed in hepatic
tissues where they primarily play a detoxification role. However, they are also
found in many other tissues such as the kidneys, intestines, brain, and
respiratory tract [12].
Cytochromes P450 were originally distinguished from other heme
proteins by their spectral features. After reduction with dithionite and the
addition of carbon monoxide gas, CYP450 enzymes demonstrate strong
5
absorption at a wavelength of 450 nm [15]. The carbon monoxide binds tightly
to the ferrous heme, causing a difference in the absorbance spectrum. This
change in absorbance is called a reduced CO difference spectrum (Figure 1.1
A). Garfinkel and Klingenberg independently observed this spectrum in the
1950s, and in 1958 Omura and Sato identified this spectrum as a
characteristic of a hemoprotein, cytochrome P450 [16-18]. Hence, the name
cytochrome P450 derives from the fact that these proteins have a heme
group, and unusual spectral properties (pigment absorbing at 450 nm).
Today, CO difference assays are used to quantitate the amount of CYP450 in
the active state.
The reason why CYP450s absorb light at 450 nm is the nature of the
atoms that interact with the heme iron. As in all heme proteins, the iron is
bonded to four nitrogens in the planar protoporphyrin IV prosthetic group
(Figure 1.2), leaving the opportunity for coordination of two additional diaxial
bonding interactions. In CYP450, the fifth proximal bond is an interaction with
a thiolate anion. This sulfur ion is provided by a conserved cysteine near the
C-terminal end of the protein. The sixth coordination position is free to bind
ligands that have entered the CYP450 active site. In a CO difference assay,
carbon monoxide is coordinating as the sixth ligand. When CYP450 is in the
normal resting state, a single water molecule occupies this position. The high
affinity of the reduced CYP450 for carbon monoxide, together with the unique
6
Figure 1.1: A reduced carbon monoxide difference spectra of CYP450 2E1:
A is a spectrum of active P450 protein, B is a spectrum of inactive P420 protein.
A
B
Figure 1.2: The heme moiety of the CYP450 enzymes with carbon monoxide
complexed to the reduced iron.
O+
CN
Fe
N!
2+
N
N
O
OH
OH
Scys
O
7
spectral properties of the CO-P450 complex were instrumental in the
discovery of this superfamily of enzymes.
Occasionally, the reduced CO difference assay can produce a maximal
absorption band at 420 nm instead of 450 nm (Figure 1.1 B). This spectral
shift to 420 nm is attributed to the protonation of the cysteine thiolate to form a
neutral thiol [19]. For catalytic activity, the cysteine must be deprotonated
throughout the entire reaction cycle. The protonation of the cysteine most
likely occurs during a change in the environment of the heme-binding pocket
allowing a proton to bond to the cysteine thiolate [20]. Changes in the protein
that facilitate protonation are currently not well understood, but may have
multiple causes including large or small scale conformational changes or
partial unfolding. These changes result in catalytically inactive protein and are
generally irreversible, though in a few cases at least some recovery of the
active CYP450 form has been demonstrated [21].
CYP450 Enzymes: Organization of Isozymes and Nomenclature
After significant scientific debate during the 1970s, it became
increasingly apparent that there are multiple CYP450s in different organisms
and within a single organism or even a single tissue [22-29]. In the
postgenomic era, we know that some organisms have as many as 323
8
cytochrome P450 enzymes [30]. As a result of this expanded recognition of
CYP450 diversity, a systematic nomenclature was devised to organize this
large superfamily of enzymes into families, subfamilies, and individual
isozymes [31].
Nebert organized the CYP450 superfamily into families based on their
structural and evolutionary correlations [32]. The families and subfamilies are
classed by their amino acid sequence identities [32].
Enzymes whose
sequences are greater than 40% identical at the amino acid level are grouped
into the same family, denoted by the initial Arabic number in the name of the
enzyme.
Enzymes whose sequences are greater than 55% identical are
grouped into the same subfamily, denoted by a capitalized alphabetic
character following the family numeral. Finally, a terminal Arabic number
identifies the individual enzymes within a subfamily. Examples of the
nomenclature used to identify a cytochrome isoenzyme are: CYP1A2,
CYP2E1, and CYP3A4. Since function usually follows overall structure, this
sequence-based nomenclature system provides a general grouping of
enzymes with relatively similar substrate selectivity. Although there are
examples known whereby CYP450 enzymes in different subfamilies and even
families metabolize the same substrate, the regio- and stereoselectivity of
metabolism is often significantly different in these cases.
9
CYP450 Structure and Active Site Topography
It is widely known that many CYP450 enzymes differ in their selectivity
for substrates and inhibitors and even when two enzymes can metabolize the
same substrate, they can exhibit widely different turnover numbers. Despite
their broad range of substrates, many general structural features are
conserved among all CYP450s as evidenced by the structures of ten
published mammalian CYP450 enzymes [33-45].
Of the membrane bound CYP450 enzymes crystallized to date, all
possess a triangular prism shape containing twelve major α-helical segments,
designated helices A-L [33] (Figure 1.3). Typically, the F thru G portion lie
orthogonal to the structurally conserved I helix that anchors the heme
cofactor. In the mammalian CYP2B4, Scott et al. has shown that the B′ to C
and F to G regions can be dramatically repositioned to create a cleft that
would allow substrates entry to the active site [46]. The long I helix extends
the length of the entire CYP structure, and has a kink in the proximity of the
heme cofactor. Embedded within the enzyme, the heme cofactor iron
protoporphyrin IX (Figure 1.2) is located between the L helix on the proximal
side and the I helix on the distal side and covalently bound to the thiolate ion
of a cysteine residue preceding the L helix. The remaining heme-binding
10
Figure 1.3: The crystal structure of CYP2A13 in a closed conformation. The SRS
regions are highlighted by color: SRS-1 (yellow), SRS-2 (orange), SRS-3 (pink),
SRS-4 (green), SRS-5 (blue), and SRS-6 (purple).
D
G
F´
G´
A´
F
B´
E
I
A
K
J
H
L
B
C
11
interactions consist of complementary hydrophobic interactions and hydrogen
bonds between protein side chains and the heme propionate groups.
CYP450s from the CYP4 family have an additional ester bond between the
heme and the apoprotein [47-49].
In cytochromes P450, the active site is proximal to the heme and is
defined by the convergence of six non-continuous stretches of amino acids
that interact with substrates called substrate recognition sites (SRS) [50]. A
short helix between the B and C helices (designated B′) composes the SRS-1
region. The SRS-2 region includes the C-terminus of the F helix. A portion of
the G helix composes the SRS-3 region. A segment of the I helix makes up
the SRS-4 region. The SRS-5 region, a β-strand following the K-helix, and
SRS-6, a β-loop near the C-terminus of the enzyme, make up the last wall of
the active site. Of these six regions, the F/G region including SRS-2 and the
B′ helix (SRS-1) seem to be the most flexible. In several crystal structures, the
B′ helix is repositioned in response to the identity of the ligand in the active
site in something of an induced-fit mode. The C-terminus of helix F may also
be involved in induced fit to various ligands in the active site, but likely also
plays a role in conformational changes required for ligand entry and exit in
association with the short subsequent helices F′ and G′, and the long G helix.
12
An evaluation of the substrate free and substrate bound structures,
both in bacterial and mammalian enzymes, indicates that an induced fit may
be common for most multifunctional CYP450s. The presence of both open
and closed conformations enables the enzyme to capture substrates and
achieve turnover by promoting catalysis in the closed conformation.
Substrate-bound crystal structures have offered constructive insights into the
ability of CYP450s to metabolize a wide variety of substrates. However,
despite the information about specific substrate-enzyme interactions, the
chemistry utilized by CYP450 enzymes to achieve the conversion of
substrates is an area of continuing research.
The Catalytic Cycle
In the face of the diversity and flexibility of the CYP450 enzymes, it is
interesting to note that they all operate similarly. An impressive characteristic
of this family of enzymes is the capability to generate a reactive oxygen
species from molecular oxygen and to incorporate that oxygen molecule into
a hydrophobic substrate. The catalytic cycle by which this occurs can be
separated into five parts: 1.) substrate binding, 2.) oxygen binding, 3.)
dioxygen scission, 4.) oxygen insertion into the substrate, and 5.) product
release (Scheme 1.1).
13
Scheme 1.1: The catalytic cycle of cytochromes P450.
14
The first step involves the reversible binding and orientation of the
substrate in the active site with the iron in its ferric state. This process often
displaces an iron-bound water molecule typically found in the resting state of
the enzyme and is responsible for some unique spectral changes. Once the
substrate has been bound, two electrons are delivered to NADPH and one of
those electrons via cytochrome P450-oxidoreductase, reduces the iron to its
ferrous form. After the first electron is delivered, the iron-substrate complex
binds molecular oxygen in the second step of the catalytic cycle. Once bound,
the molecular oxygen must be reduced to split the double bond between the
two oxygen molecules. The third step of cleaving the oxygen molecule
requires the rearrangement of the peroxo-radical complex to form the
superoxide anion. A second electron from either cytochrome b5 or cytochrome
P450-oxidoreductase reduces the complex to a peroxoanion intermediate.
The terminal oxygen atom will then bind two hydrogen atoms to form water,
leaving a single oxygen atom bound to the iron, probably in a perferryl
complex. During the fourth step, it is thought that the substrate is activated by
either removing hydrogen (hydrogen abstraction) or an electron (e.g. from
nitrogen atoms) from the substrate molecule, leaving the carbon as a reactive
radical. The activated substrate is now free to react with the heme-bound
activated oxygen. Once the substrate has been converted to a metabolite, it
has transformed enough that it is no longer energetically favorable for it to
15
linger in the active site. Therefore, during the fifth and final step the metabolite
is released and the enzyme is free to repeat the process.
CYP450 2 Family: Substrate Overlap and Diversity
In humans, the CYP2 family is the single largest family comprising
approximately 28 percent of the CYP450s. The CYP2 family consists of
eleven subfamilies containing sixteen isozymes. Approximately 50% of all the
drugs currently on the market are metabolized by CYP450s in the CYP2
family (Figure 1.4) [51]. In particular, this family is known for its broad (and in
many cases overlapping) range of substrate specificities.
Members of the CYP2 family are flexible enough to metabolize many
potential toxins (e.g. drugs, herbs, and pollutants). The levels of these
xenobiotic-metabolizing CYP450s may vary considerably, in contrast to the
fairly consistent levels of most CYP450s that have crucial endogenous roles.
Table 1.1 lists the sixteen human CYP2 family isozymes, along with available
knowledge about sites of tissue expression, subcellular localization, a typical
reaction, and known inducers. As this work is concerned with CYP450
enzymes from the 2A and 2E subfamilies, these enzymes will be discussed
further.
16
Figure 1.4: Major CYP450 enzymes responsible for phase I metabolism. The
percentage of phase I metabolism of drugs that each enzyme contributes to is
demonstrated by the relative size of each slice of the chart [51, 52].
17
Table 1.1: The human CYP2 family of CYP450s and their known locations,
reactions, and inducers [12]. Abbreviation ER designates endoplasmic reticulum.
CYP450
Tissue Sites
Subcellular
Localization
Typical
Reaction
Known
Inducers
2A6
Liver, lung, and
some
extrahepatic
sites
ER
Coumarin
7-hydroxylation
Phenobarbital
Barbiturates
Dexamethasone
Rifampin
ER
ER
2
2
Activation of NNK
2
ER
(S)-Mephenytoin
N-demethylation
Taxol
6α-hydroxylation
Tobutamine methyl
hydroxylation
Phenobarbital
Nicotine
Phenobarbital
Rifampin
Phenobarbital
Rifampin
Dexamethasone
Phenobarbital
Phenobarbital
Refampin
Nicotine
2A7
2A13
2B6
Respiratory
Tract
Liver, lung
2C8
Liver
ER
2C9
Liver
ER
2C18
2C19
Liver
Liver
ER
ER
2D6
Liver
ER1
2E1
ER
2F1
Liver, lung,
other tissues
Lung
2J2
Lung
ER
2R1
2S1
2U1
2W1
2
2
2
2
ER
ER
ER
2
2
2
2
2
2
Lung
2
2
1
2
ER
2
(S)-Mephenytoin
4′-hydroxylation
Debrisoquine
4-hydroxylation
Chlorzoxazone
6-hydroxylation
3-Methylindole
activation
Arachidonic
acid oxidations
Acetone, Ethanol,
Isoniazid, Nicotine
2
2
Mainly ER, some detected in mitochondria.
Currently unknown.
18
CYP450 2A6
CYP2A6 comprises from <0.2-13% of the total liver CYP450 content,
with levels differing dramatically between individuals [53]. CYP2A6 is clinically
inducible by anticonvulsants such as phenobarbital and the antibacterial
rifampicin [54, 55].
The CYP2A substrate selectivity exhibits some amount of overlap with
enzymes of the CYP2B and CYP2E subfamilies [56]. Many of the CYP2A6
substrates contain a ketone functional group and are relatively polar mediumto-low molecular weight compounds with hydrogen bond acceptor atoms
relatively close to the favored site of metabolism (Figure 1.5). CYP2A6 is
known to metabolize coumarin to 7-hydroxycoumarin, and this activity has
been used as a marker substrate for the enzyme for several years [57, 58].
Because of this, the 7-hydroxylation of coumarin has been used as an in vivo
diagnostic assay for this isozyme [59-61]. CYP2A6 is responsible for the
metabolic conversion of nicotine to cotinine, and the further hydroxylation of
cotinine [62-64]. In addition, CYP2A6 catalyzes the 2′-hydroxylation of
nicotine to a lung procarcinogen [65]. Methoxalen (an antipsoriasis agent) and
flavenoids in grapefruit juice are potent mechanism-based inhibitors of
CYP2A6 [66, 67]. Imidazoles are also known to be weak inhibitors of this
enzyme [68]. CYP2A6 plays a toxicological role, in that it metabolizes
carcinogens including aflatoxins, 1,3 butadiene, and nitrosamines [69-72].
19
Figure 1.5: Structures of representative CYP2A6 substrates.
Arrow denotes preferred site of metabolism.
O
O
N
coumarin
nicotine
O
N
N
N
cotinine
fadrozole
Cl
CN
Cl
F
O
Cl
O
OH
O
F
methoxyflurane
O
losigamone
20
CYP450 2A13
CYP2A13 has substrate specificity similar to that of CYP2A6 (Figure
1.6), because of the high amino acid identity between the two isozymes
(93.5%). Therefore, it is not surprising that both are important enzymes in the
metabolism of nicotine and cotinine [73]. CYP2A13 has been identified as the
more efficient enzyme in nicotine metabolism, but because of its primary
expression in the respiratory tract, the bulk of systemic nicotine is typically
metabolized by the liver CYP2A6. CYP2A13 is also involved in the metabolic
activation of 4-(methylnitrosamino)-1-(3-pyridyl)-1butanone (NNK), found in
cigarettes and can also be produced endogenously from the metabolism of
nicotine [74-77]. NNK metabolic activation requires the hydroxylation at one of
the two α carbons to the N-nitroso group, leading to the formation of reactive
intermediates that can form either DNA adducts or stable metabolites [76].
This is of particular interest with regard to tobacco-related cancer due to the
localization of CYP2A13. As mentioned, this enzyme is primarily expressed in
the respiratory tract, with the highest level in the nasal mucosa, followed by
the lung and trachea.
21
Figure 1.6: Structures of representative CYP2A13 substrates.
Arrow denotes preferred site of metabolism and wavy line
represents dealkylation. Arrow in the center of a bond represents
epoxidation.
H
N
O
O
styrene
phenacetin
N
O
N
cotinine
naphthalene
O
N
NH
N
O
N
N
N
nicotine
theophylline
O
O
N
N
N
4-(methylnitrosamino)-1-(3-pyridyl)-1butanone
22
CYP450 2E1
CYP2E1 comprises approximately 10 percent of human liver CYP450
and is relatively unusual in that it oxidizes small industrial solvents, ranging
from pyridine to ethanol, acetone, and other small ketones [53, 78]. Ethanol
and acetone are also known as strong inducers of this isoform [79]. Many of
its substrates are water-soluble and often implicated in toxicity (Figure 1.7).
CYP2E1 is involved in the activation of several carcinogens and other toxic
chemicals (e.g. benzene, dialkylnitrosamines, and halothanes) [80, 81].
CYP2E1 is also known to metabolize a moderately small number of
pharmaceutical agents, including chlorzoxazone and paracetamol [51].
The generation of oxygen radicals and other reactive oxygen species
(ROS) has also been associated with CYP2E1. The oxygen activation
process, mentioned previously, is the generally accepted catalytic progression
to generate the active oxidant and substrate turnover. However, some
enzymes (e.g. CYP2E1) have been observed to release reactive oxygen
species in the form of superoxide, peroxy radical, and/or hydrogen peroxide
[82-84]. When this occurs the reaction is said to be “uncoupled.” This
uncoupling can lead to an inefficient turnover of substrate and/or cause
damage to neighboring cellular components. The creation of ROS is likely due
to a ”short circuit” in the enzymatic proton-transfer system involved in the
23
Figure 1.7: Structures of representative CYP2E1 substrates.
Arrows represent preferred site of metabolism.
NO2
O
HO
N
H
paracetamol (acetaminophen)
OH
p-nitrophenol
NH2
O
N
N
Diethylnitrosamine
aniline
benzene
O
OH
Cl
N
OH
ethanol
chlorzoxazone
24
transformation of the enzyme-bound oxygen intermediates to the final active
oxidant.
Enzyme Kinetics
Enzymes are biological macromolecules that enhance the rates of
chemical transformations [85]. Enzymes provide a micro-environment that
provides optimal conditions necessary to increase the rate of reaction.
Cytochromes P450 are unusual in the diversity of the substrates metabolized
and the metabolites generated by a single enzyme. Most enzymes are
uniquely selective in the reactants and products of the chemical reactions
they catalyze.
Enzyme kinetics is the study of the rate at which an enzyme catalyzes
a metabolic transformation [86]. The rate at which enzymes operate is
influenced by several factors, including substrate concentration, the presence
of inhibitors, and environmental parameters such as the temperature and the
pH. There are multiple techniques for monitoring the rates of enzyme
reactions by measuring the disappearance of substrate or the appearance of
product.
equation
In 1913, Leonor Michaelis and Maud Menten introduced the
Vo =
V max[S]
Km + [S]
[87].
The
equation
describes
the
hyperbolic
!
25
dependence of Vo on substrate concentration, which occurs for most
enzymes. Figure 1.8 illustrates the relationships between this equation and
enzyme behavior at low and high substrate concentrations. At low
concentrations, the substrate term in the denominator of the MichaelisMenten equation becomes negligible. The equation will now simplify to
Vo=(Vmax [S])/Km and Vo demonstrates a linear dependence on the substrate
concentration. At high concentrations, the Km term in the denominator
becomes negligible, and the equation simplifies to Vo=Vmax. This is consistent
with the plateau observed at high substrate concentrations. Accordingly, the
Michaelis-Menten equation is consistent with the dependence of Vo on [S],
and the shape of the curve is defined by the terms Vmax/Km at low substrate
concentrations and Vmax at high concentrations.
Understanding the three-dimensional structures of enzymes provides
insights into the mechanism of their action. The importance of structural
information is complemented by classical protein chemistry and site-directed
mutagenesis. These methods aid in examining the importance of individual
amino acids in enzyme structure and function. Determining the rate of
reaction and how it changes is a core technique for studying the mechanisms
of an enzyme-catalyzed reaction.
26
Figure 1.8: A plot of substrate concentration versus the initial velocity
of an enzyme-catalyzed reaction.
Initial velocity, Vo (e.g. µM/min)
Vo =
Vmax[S]
Km
Vo = Vmax
!
1/2 Vmax
Km
Substrate Concentration (e.g. µM)
27
References
1
Editors of the American Heritage Diction. (2004) The American Heritage
Stedman's Medical Dictionary. Houghton Mifflin Company
2
Silverman, R. B. (2004) The Organic Chemistry of Drug Design and Drug
Action. Elsevier Academic Press, San Diego
3
Yengi, L. G., Leung, L. and Kao, J. (2007) The evolving role of drug
metabolism in drug discovery and development. Pharm Res 24, 842-858
4
Coleman, M. D. (2005) Human Drug Metabolism: An Introduction. John Wiley
& Sons, Ltd, West Sussex
5
Block, J. H. and Beale, J.M. (2004) Wilson and Gisvold's Textbook of Organic
Medicinal and Pharmaceutical Chemistry. Lippincott Williams & Wilkins,
Baltimore
6
Williams, R. T. (1959) Detoxification Mechanisms. John Wiley & Sons, New
York
7
Drayer, D. E. (1982) Pharmacologically active metabolites of drugs and other
foreign compounds. Clinical, pharmacological, therapeutic and toxicological
considerations. Drugs 24, 519-542
8
Woolf, T. F. (1999) Handbook of Drug Metabolism. Marcel Dekker, New York
9
Schenkman, J. B. (1999) The fate of xenobiotics in the body: Enzymes of
metabolism. NATO Adv. Sci. Inst. Ser. A Life Sci. 303, 1-20
10
Lewis, D. F. V. (2001) Guide to Cytochromes P450 Structure and Function.
Taylor and Francis Inc., London
11
Waterman, M. R. (1996) Introduction: transcription and regulation of activities
of cytochromes P450 metabolizing endogenous substrates. Faseb J 10, 1455
12
Ortiz de Montellano, P. R. (2004) Cytochrome P450: Structure, Mechanism,
and Biochemistry. Springer-Verlag New York, LLC, New York
13
Adas, F., Salaun, J. P., Berthou, F., Picart, D., Simon, B. and Amet, Y. (1999)
Requirement for omega and (omega;-1)-hydroxylations of fatty acids by
human cytochromes P450 2E1 and 4A11. J Lipid Res 40, 1990-1997
28
14
Sono, M., Roach, M. P., Coulter, E. D. and Dawson, J. H. (1996) HemeContaining Oxygenases. Chem Rev 96, 2841-2888
15
Omura, T. and Sato, R. (1964) The Carbon Monoxide-Binding Pigment of
Liver Microsomes. I. Evidence for Its Hemoprotein Nature. J Biol Chem 239,
2370-2378
16
Garfinkel, D. (1958) Studies on pig liver microsomes. I. Enzymic and pigment
composition of different microsomal fractions. Arch Biochem Biophys 77, 493509
17
Klingenberg, M. (1958) Pigments of rat liver microsomes. Arch Biochem
Biophys 75, 376-386
18
Omura, T. and Sato, R. (1962) A new cytochrome in liver microsomes. J Biol
Chem 237, 1375-1376
19
Perera, R., Sono, M., Sigman, J. A., Pfister, T. D., Lu, Y. and Dawson, J. H.
(2003) Neutral thiol as a proximal ligand to ferrous heme iron: implications for
heme proteins that lose cysteine thiolate ligation on reduction. Proc Natl Acad
Sci U S A 100, 3641-3646
20
Anzenbacherova, E., Hudecek, J., Murgida, D., Hildebrandt, P., Marchal, S.,
Lange, R. and Anzenbacher, P. (2005) Active sites of two orthologous
cytochromes P450 2E1: differences revealed by spectroscopic methods.
Biochem Biophys Res Commun 338, 477-482
21
Ho, W. W., Li, H., Nishida, C. R., de Montellano, P. R. and Poulos, T. L.
(2008)
Crystal
structure
and
properties
of
CYP231A2
from
the
thermoacidophilic archaeon Picrophilus torridus. Biochemistry 47, 2071-2079
22
Bandiera, S., Ryan, D. E., Levin, W. and Thomas, P. E. (1985) Evidence for a
family of four immunochemically related isozymes of cytochrome P-450
purified from untreated rats. Arch Biochem Biophys 240, 478-482
23
Guengerich, F. P. (1977) Separation and purification of multiple forms of
microsomal cytochrome P-450. Activities of different forms of cytochrome P450 towards several compounds of environmental interest. J Biol Chem 252,
3970-3979
29
24
Guengerich, F. P. (1979) Isolation and purification of cytochrome P-450, and
the existence of multiple forms. Pharmacol Ther 6, 99-121
25
Haugen, D. A., van der Hoeven, T. A. and Coon, M. J. (1975) Purified liver
microsomal cytochrome P-450. Separation and characterization of multiple
forms. J Biol Chem 250, 3567-3570
26
Huang, M. T., West, S. B. and Lu, A. Y. (1976) Separation, purification, and
properties of multiple forms of cytochrome P-450 from the liver microsomes of
phenobarbital-treated mice. J Biol Chem 251, 4659-4665
27
Ryan, D. E., Iida, S., Wood, A. W., Thomas, P. E., Lieber, C. S. and Levin, W.
(1984) Characterization of three highly purified cytochromes P-450 from
hepatic microsomes of adult male rats. J Biol Chem 259, 1239-1250
28
Ryan, D. E., Thomas, P. E., Parkinson, A., Reik, L. M., Wood, A. W. and
Levin, W. (1984) Characterization and regulation of rat hepatic microsomal
cytochrome P-450 isozymes. Ann N Y Acad Sci 435, 73-85
29
Porter, T. D. (2004) Jud Coon: 35 years of p450 research, a synopsis of p450
history. Drug Metab Dispos 32, 1-6
30
Nelson, D. (2004) Numbers of putatively functional full-length CYP genes in
54 whole eukaryotic genomes.
31
Nebert, D. W., Nelson, D. R., Adesnik, M., Coon, M. J., Estabrook, R. W.,
Gonzalez, F. J., Guengerich, F. P., Gunsalus, I. C., Johnson, E. F., Kemper,
B. and et al. (1989) The P450 superfamily: updated listing of all genes and
recommended nomenclature for the chromosomal loci. DNA 8, 1-13
32
Nebert, D. W., Nelson, D. R., Coon, M. J., Estabrook, R. W., Feyereisen, R.,
Fujii-Kuriyama, Y., Gonzalez, F. J., Guengerich, F. P., Gunsalus, I. C.,
Johnson, E. F. and et al. (1991) The P450 superfamily: update on new
sequences, gene mapping, and recommended nomenclature. DNA Cell Biol
10, 1-14
33
Otyepka, M., Skopalik, J., Anzenbacherova, E. and Anzenbacher, P. (2007)
What common structural features and variations of mammalian P450s are
known to date? Biochim Biophys Acta 1770, 376-389
30
34
Strushkevich, N. V., Min, J., Loppnau, P., Tempel, W., Arrowsmith, C.H.,
Edwards, A.M., Sundstrom, M., Weigelt, J., Bochkarev, A., Plotnikov, A.N.,
Park, H., Structural Genomics Consortium (SGC) (2008) Structural analysis
of CYP2R1 in complex with vitamin D3., Structural Genomics
35
Yano, J. K., Hsu, M. H., Griffin, K. J., Stout, C. D. and Johnson, E. F. (2005)
Structures of human microsomal cytochrome P450 2A6 complexed with
coumarin and methoxsalen. Nat Struct Mol Biol 12, 822-823
36
Scott, E. E., He, Y. A., Wester, M. R., White, M. A., Chin, C. C., Halpert, J. R.,
Johnson, E. F. and Stout, C. D. (2003) An open conformation of mammalian
cytochrome P450 2B4 at 1.6-A resolution. Proc Natl Acad Sci U S A 100,
13196-13201
37
Williams, P. A., Cosme, J., Sridhar, V., Johnson, E. F. and McRee, D. E.
(2000) Mammalian microsomal cytochrome P450 monooxygenase: structural
adaptations for membrane binding and functional diversity. Mol Cell 5, 121131
38
Wester, M. R., Yano, J. K., Schoch, G. A., Yang, C., Griffin, K. J., Stout, C. D.
and Johnson, E. F. (2004) The structure of human cytochrome P450 2C9
complexed with flurbiprofen at 2.0-A resolution. J Biol Chem 279, 3563035637
39
Williams, P. A., Cosme, J., Ward, A., Angove, H. C., Matak Vinkovic, D. and
Jhoti, H. (2003) Crystal structure of human cytochrome P450 2C9 with bound
warfarin. Nature 424, 464-468
40
Rowland, P., Blaney, F. E., Smyth, M. G., Jones, J. J., Leydon, V. R.,
Oxbrow, A. K., Lewis, C. J., Tennant, M. G., Modi, S., Eggleston, D. S.,
Chenery, R. J. and Bridges, A. M. (2006) Crystal structure of human
cytochrome P450 2D6. J Biol Chem 281, 7614-7622
41
Williams, P. A., Cosme, J., Vinkovic, D. M., Ward, A., Angove, H. C., Day, P.
J., Vonrhein, C., Tickle, I. J. and Jhoti, H. (2004) Crystal structures of human
cytochrome P450 3A4 bound to metyrapone and progesterone. Science 305,
683-686
31
42
Yano, J. K., Wester, M. R., Schoch, G. A., Griffin, K. J., Stout, C. D. and
Johnson, E. F. (2004) The structure of human microsomal cytochrome P450
3A4 determined by X-ray crystallography to 2.05-A resolution. J Biol Chem
279, 38091-38094
43
Smith, B. D., Sanders, J. L., Porubsky, P. R., Lushington, G. H., Stout, C. D.
and Scott, E. E. (2007) Structure of the human lung cytochrome P450 2A13.
J Biol Chem 282, 17306-17313
44
Sansen, S., Yano, J. K., Reynald, R. L., Schoch, G. A., Griffin, K. J., Stout, C.
D. and Johnson, E. F. (2007) Adaptations for the oxidation of polycyclic
aromatic hydrocarbons exhibited by the structure of human P450 1A2. J Biol
Chem 282, 14348-14355
45
Schoch, G. A., Yano, J. K., Wester, M. R., Griffin, K. J., Stout, C. D. and
Johnson, E. F. (2004) Structure of human microsomal cytochrome P450 2C8.
Evidence for a peripheral fatty acid binding site. J Biol Chem 279, 9497-9503
46
Scott, E. E., White, M. A., He, Y. A., Johnson, E. F., Stout, C. D. and Halpert,
J. R. (2004) Structure of mammalian cytochrome P450 2B4 complexed with
4-(4-chlorophenyl)imidazole at 1.9-A resolution: insight into the range of P450
conformations and the coordination of redox partner binding. J Biol Chem
279, 27294-27301
47
LeBrun, L. A., Xu, F., Kroetz, D. L. and Ortiz de Montellano, P. R. (2002)
Covalent attachment of the heme prosthetic group in the CYP4F cytochrome
P450 family. Biochemistry 41, 5931-5937
48
LeBrun, L. A., Hoch, U. and Ortiz de Montellano, P. R. (2002) Autocatalytic
mechanism and consequences of covalent heme attachment in the
cytochrome P4504A family. J Biol Chem 277, 12755-12761
49
Hoch, U. and Ortiz De Montellano, P. R. (2001) Covalently linked heme in
cytochrome p4504a fatty acid hydroxylases. J Biol Chem 276, 11339-11346
50
Gotoh, O. (1992) Substrate recognition sites in cytochrome P450 family 2
(CYP2) proteins inferred from comparative analyses of amino acid and
coding nucleotide sequences. J Biol Chem 267, 83-90
32
51
Rendic, S. and Di Carlo, F. J. (1997) Human cytochrome P450 enzymes: a
status report summarizing their reactions, substrates, inducers, and inhibitors.
Drug Metab Rev 29, 413-580
52
Evans, W. E. and Relling, M. V. (1999) Pharmacogenomics: translating
functional genomics into rational therapeutics. Science 286, 487-491
53
Shimada, T., Yamazaki, H., Mimura, M., Inui, Y. and Guengerich, F. P. (1994)
Interindividual variations in human liver cytochrome P-450 enzymes involved
in the oxidation of drugs, carcinogens and toxic chemicals: studies with liver
microsomes of 30 Japanese and 30 Caucasians. J Pharmacol Exp Ther 270,
414-423
54
Rae, J. M., Johnson, M. D., Lippman, M. E. and Flockhart, D. A. (2001)
Rifampin is a selective, pleiotropic inducer of drug metabolism genes in
human hepatocytes: studies with cDNA and oligonucleotide expression
arrays. J Pharmacol Exp Ther 299, 849-857
55
Donato, M. T., Viitala, P., Rodriguez-Antona, C., Lindfors, A., Castell, J. V.,
Raunio, H., Gomez-Lechon, M. J. and Pelkonen, O. (2000) CYP2A5/CYP2A6
expression in mouse and human hepatocytes treated with various in vivo
inducers. Drug Metab Dispos 28, 1321-1326
56
Lewis, D. F., Dickins, M., Lake, B. G., Eddershaw, P. J., Tarbit, M. H. and
Goldfarb, P. S. (1999) Molecular modelling of the human cytochrome P450
isoform CYP2A6 and investigations of CYP2A substrate selectivity.
Toxicology 133, 1-33
57
Yun, C. H., Shimada, T. and Guengerich, F. P. (1991) Purification and
characterization of human liver microsomal cytochrome P-450 2A6. Mol
Pharmacol 40, 679-685
58
Yamano, S., Tatsuno, J. and Gonzalez, F. J. (1990) The CYP2A3 gene
product catalyzes coumarin 7-hydroxylation in human liver microsomes.
Biochemistry 29, 1322-1329
59
Daly, A. K., Cholerton, S., Gregory, W. and Idle, J. R. (1993) Metabolic
polymorphisms. Pharmacol Ther 57, 129-160
33
60
Cholerton, S., Idle, M. E., Vas, A., Gonzalez, F. J. and Idle, J. R. (1992)
Comparison of a novel thin-layer chromatographic-fluorescence detection
method with a spectrofluorometric method for the determination of 7hydroxycoumarin in human urine. J Chromatogr 575, 325-330
61
Rautio, A., Kraul, H., Kojo, A., Salmela, E. and Pelkonen, O. (1992)
Interindividual variability of coumarin 7-hydroxylation in healthy volunteers.
Pharmacogenetics 2, 227-233
62
Nakajima, M., Yamamoto, T., Nunoya, K., Yokoi, T., Nagashima, K., Inoue,
K., Funae, Y., Shimada, N., Kamataki, T. and Kuroiwa, Y. (1996) Role of
human cytochrome P4502A6 in C-oxidation of nicotine. Drug Metab Dispos
24, 1212-1217
63
Messina, E. S., Tyndale, R. F. and Sellers, E. M. (1997) A major role for
CYP2A6 in nicotine C-oxidation by human liver microsomes. J Pharmacol
Exp Ther 282, 1608-1614
64
Yamazaki, H., Inoue, K., Hashimoto, M. and Shimada, T. (1999) Roles of
CYP2A6 and CYP2B6 in nicotine C-oxidation by human liver microsomes.
Arch Toxicol 73, 65-70
65
Nakajima, M., Yamamoto, T., Nunoya, K., Yokoi, T., Nagashima, K., Inoue,
K., Funae, Y., Shimada, N., Kamataki, T. and Kuroiwa, Y. (1996)
Characterization of CYP2A6 involved in 3'-hydroxylation of cotinine in human
liver microsomes. J Pharmacol Exp Ther 277, 1010-1015
66
Hukkanen, J., Jacob, P., 3rd and Benowitz, N. L. (2006) Effect of grapefruit
juice on cytochrome P450 2A6 and nicotine renal clearance. Clin Pharmacol
Ther 80, 522-530
67
Zhang, W., Kilicarslan, T., Tyndale, R. F. and Sellers, E. M. (2001) Evaluation
of methoxsalen, tranylcypromine, and tryptamine as specific and selective
CYP2A6 inhibitors in vitro. Drug Metab Dispos 29, 897-902
68
Zhang, W., Ramamoorthy, Y., Kilicarslan, T., Nolte, H., Tyndale, R. F. and
Sellers, E. M. (2002) Inhibition of cytochromes P450 by antifungal imidazole
derivatives. Drug Metab Dispos 30, 314-318
34
69
Yamazaki, H., Inui, Y., Yun, C. H., Guengerich, F. P. and Shimada, T. (1992)
Cytochrome P450 2E1 and 2A6 enzymes as major catalysts for metabolic
activation of N-nitrosodialkylamines and tobacco-related nitrosamines in
human liver microsomes. Carcinogenesis 13, 1789-1794
70
Smith, T. J., Guo, Z., Gonzalez, F. J., Guengerich, F. P., Stoner, G. D. and
Yang, C. S. (1992) Metabolism of 4-(methylnitrosamino)-1-(3-pyridyl)-1butanone in human lung and liver microsomes and cytochromes P-450
expressed in hepatoma cells. Cancer Res 52, 1757-1763
71
Mace, K., Aguilar, F., Wang, J. S., Vautravers, P., Gomez-Lechon, M.,
Gonzalez, F. J., Groopman, J., Harris, C. C. and Pfeifer, A. M. (1997)
Aflatoxin B1-induced DNA adduct formation and p53 mutations in CYP450expressing human liver cell lines. Carcinogenesis 18, 1291-1297
72
Duescher, R. J. and Elfarra, A. A. (1994) Human liver microsomes are
efficient catalysts of 1,3-butadiene oxidation: evidence for major roles by
cytochromes P450 2A6 and 2E1. Arch Biochem Biophys 311, 342-349
73
Bao, Z., He, X. Y., Ding, X., Prabhu, S. and Hong, J. Y. (2005) Metabolism of
nicotine and cotinine by human cytochrome P450 2A13. Drug Metab Dispos
33, 258-261
74
Su, T., Bao, Z., Zhang, Q. Y., Smith, T. J., Hong, J. Y. and Ding, X. (2000)
Human cytochrome P450 CYP2A13: predominant expression in the
respiratory tract and its high efficiency metabolic activation of a tobaccospecific carcinogen, 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanone. Cancer
Res 60, 5074-5079
75
Zhang, X., D'Agostino, J., Wu, H., Zhang, Q. Y., von Weymarn, L., Murphy, S.
E. and Ding, X. (2007) CYP2A13: variable expression and role in human lung
microsomal metabolic activation of the tobacco-specific carcinogen 4(methylnitrosamino)-1-(3-pyridyl)-1-butanone. J Pharmacol Exp Ther 323,
570-578
76
Hecht, S. S. (1998) Biochemistry, biology, and carcinogenicity of tobaccospecific N-nitrosamines. Chem Res Toxicol 11, 559-603
35
77
Hecht, S. S., Hochalter, J. B., Villalta, P. W. and Murphy, S. E. (2000) 2'Hydroxylation of nicotine by cytochrome P450 2A6 and human liver
microsomes: formation of a lung carcinogen precursor. Proc Natl Acad Sci U
S A 97, 12493-12497
78
Ronis, M. J. J., Lindros, K. O. and Ingelman-Sundberg, M. (1987) The CYP2E
subfamily, in: Cytochromes P450 - metabolic and toxicological aspects. CRC
Press, Boca Raton, FL
79
Thomas, P. E., Bandiera, S., Maines, S. L., Ryan, D. E. and Levin, W. (1987)
Regulation of cytochrome P-450j, a high-affinity N-nitrosodimethylamine
demethylase, in rat hepatic microsomes. Biochemistry 26, 2280-2289
80
Elfarra, A. A. (1993) Aliphatic halogenated hydrocarbons, in: Toxicology of
the Kidney. J.B. Hook and R.S. Goldstein, eds, Raven, New York
81
Lau, S. S. and Monks, T. J. (1993) Nephrotoxicity of bromobenzene: the role
of quinonethioethers, in: Toxicology of the Kidney. J.B. Hook and R.S.
Goldstein, eds, Raven, New York
82
White, R. E., Sligar, S. G. and Coon, M. J. (1980) Modes of oxygen activation
by cytochrome P-450. Dev. Biochem. 13, 307-314
83
McCarthy, M. B. and White, R. E. (1983) Functional differences between
peroxidase compound I and the cytochrome P-450 reactive oxygen
intermediate. J Biol Chem 258, 9153-9158
84
Perret, A. and Pompon, D. (1998) Electron shuttle between membrane-bound
cytochrome P450 3A4 and b5 rules uncoupling mechanisms. Biochemistry
37, 11412-11424
85
Matthews, J. C. (1993) Fundamentals of Receptor, Enzyme, and Transport
Kinetics. CRC Press, Inc., Boca Raton
86
Nelson, D. L. C., M.M. (2005) Lehninger Principles of Biochemsitry. W.H.
Freeman and Company, New York
87
Michaelis, L. M., Maud (1913) Die Kinetik der Invertinwirkung. Biochemistry
49, 333-369
36
Chapter 2
Project Goals, Hypothesis, and Design
Project Goals and Hypothesis
The cytochrome P450 (CYP450) superfamily of enzymes contributes to
the majority of the phase I metabolism of xenobiotics in the human body [1].
They participate in the metabolism of a greater part of the drugs found in
present clinical use and have been linked to the activation of carcinogens and
other toxins [2]. The CYP2 family, in particular, is known for its extensive
phase I metabolism of numerous xenobiotic compounds [1].
The specificity of the metabolites formed by each CYP450 isozyme
implies a set of fixed substrate-enzyme interactions, whereas the diversity of
substrates metabolized implies a flexible binding site. Understanding the
reasons for the specific, yet adaptable, metabolism by CYP450s is often
restricted by the lack of structural information for these enzymes [3].
The goal of this project is to explain the structural foundation for the
substrate specificities of the human 2A and 2E cytochrome P450 subfamilies,
which are 47.5% identical and 66.5% similar, when comparing the isoforms
2A13 and 2E1. As described in Chapter 1, CYP450 2A6, 2A13, and 2E1
37
metabolize both common and unique substrates. Therefore, it is believed that
only a small number of amino acid residues are responsible for the substrateenzyme interactions that differentiate the metabolic abilities of these two
enzymes. Identifying these interactions could help predict the metabolism of
drugs, procarcinogens, and other xenobiotics.
Our strategy is to target identification of the residues that distinguish
2E and 2A activity by exchanging the dissimilar single amino acid residues
between CYP450 2E and 2A subfamilies. This was accomplished within the
highly variable substrate recognition sites 1 and 2. To identify the amino acids
responsible for the functional differences, known 2E1 substrates were used to
determine the effects of these mutations on metabolism.
Project Design
Based on a sequence alignment comparison of the CYP2 family to the
Pseudomonas putida P450 101A (P450cam) whose substrate-binding residues
were identified by x-ray crystallography, Gotoh [4] proposed six protein
regions that interact with substrates called substrate recognition sites (SRS).
These SRSs have been generally validated by site-directed mutagenesis,
ligand-binding, enzymatic, and structural studies [5-9]. Because no structure
of CYP2E1 was available at the beginning of our study, the amino acid
38
residues in these six SRS regions were compared between the 2E and 2A
subfamilies by sequence alignment. Because substrate recognition sites 1
and 2 are highly variable between the 2A and 2E enzymes, and because of
their implications in induced fit to substrates, we focused our attention on the
dissimilar active site residues between CYP450 2E1, 2A13, and 2A6 in these
two regions.
To identify the dissimilar amino acid residues between the two CYP450
subfamilies at substrate recognition sites 1 and 2, a sequence alignment was
used. The CLC Free Workbench 4 was used to align the protein sequences
by utilizing the Dayhoff matrix. The amino acids were chosen for mutations if
they differed substantially between the two CYP450 subfamilies based on
their chemical nature (e.g., size, charge, polarity, and flexibility). For example,
glutamic acid and aspartic acid, both of which contain a carboxylic acid side
chain, would not be considered significantly different. However, a change
from a proline, a rigid “helix breaker”, to a flexible glycine would be considered
a significant difference. Figure 2.1 is the amino acid sequence alignment of
CYP450 2E1, 2A13, and 2A6. After comparison of the three sequences, 12
single amino acid mutations were chosen: CYP450 2E1 L103Q, P104A,
A105T, H107D, H109F, D111G, R112Y, W122A, P213S, W214T, L215G, and
P222S.
39
Figure 2.1: An amino acid sequence alignment of CYP450 2E1, 2A13, and 2A6.
Colored font indicates differing residue,  a 2 amino acid variance,  a 3 amino
acid variance,  the SRS-1 region, and  the SRS-2 region.
40
It is also conceivable that it is not ligand interaction with one amino
acid alone that determines substrate binding and metabolism, but the
influence of an entire face of a binding pocket that alters the activity between
enzymes. To explore this possibility, the entire sequence of the SRS-1 region
of CYP2E1 was exchanged with the SRS-1 region of CYP2A13.
Each single amino acid mutations and the SRS-1 exchange, was
engineered into a modified version of the CYP2E1 gene designed to facilitate
expression and purification of the enzyme. The full-length CYP2E1 DNA
sequence was altered by the truncation of the codons corresponding to the Nterminal transmembrane helix of the protein (Δ2–22), modification of the
codons for 23WRQVHSSWN31 to 23AKKTSSKGK31, and the addition of codons
for four sequential histidine residues at the C-terminal end of the protein.
Each mutagenesis reaction was carried out using the QuikChange® II
site-directed mutagenesis strategy. This method was chosen because it is a
highly accurate and rapid method that does not require the use of ssDNA. It
can be used to rapidly modify the DNA sequence to exchange, insert, or
delete one or more amino acids with 80% efficiency or greater, according to
the manufacturer (Stratagene). The mutated DNA plasmids were transformed
into and expressed in E. coli cells. Each mutant protein was purified by FPLC
using two columns, a NiNTA metal-affinity column and a CM ion-exchange
41
column. To identify the key amino acid residues responsible for the
differences between CYP450 2E1, 2A13, and 2A6 substrate recognition, the
2-hydroxylation of p-nitrophenol and 6-hydroxylation of chlorzoxazone (both
marker substrates for CYP2E1 [10-14]) were utilized to characterize resulting
changes in enzymatic activity.
References
1
Ortiz de Montellano, P. R. (2004) Cytochrome P450: Structure, Mechanism,
and Biochemistry. Springer-Verlag New York, LLC, New York
2
Lewis, D. F. V. (2001) Guide to Cytochromes P450 Structure and Function.
Taylor and Francis Inc., London
3
Coleman, M. D. (2005) Human Drug Metabolism: An Introduction. John Wiley
& Sons, Ltd, West Sussex
4
Gotoh, O. (1992) Substrate recognition sites in cytochrome P450 family 2
(CYP2) proteins inferred from comparative analyses of amino acid and
coding nucleotide sequences. J Biol Chem 267, 83-90
5
Kronbach, T., Larabee, T. M. and Johnson, E. F. (1989) Hybrid cytochromes
P-450 identify a substrate binding domain in P-450IIC5 and P-450IIC4. Proc
Natl Acad Sci U S A 86, 8262-8265
6
Uno, T. and Imai, Y. (1989) Identification of regions functioning in substrate
interaction of rabbit liver cytochrome P-450 (laurate (omega-1)-hydroxylase).
J Biochem 106, 569-574
7
Aoyama, T., Korzekwa, K., Nagata, K., Adesnik, M., Reiss, A., Lapenson, D.
P., Gillette, J., Gelboin, H. V., Waxman, D. J. and Gonzalez, F. J. (1989)
Sequence requirements for cytochrome P-450IIB1 catalytic activity. Alteration
of the stereospecificity and regioselectivity of steroid hydroxylation by a
42
simultaneous
change
of
two
hydrophobic
amino
acid
residues
to
phenylalanine. J Biol Chem 264, 21327-21333
8
Poulos, T. L., Finzel, B. C. and Howard, A. J. (1987) High-resolution crystal
structure of cytochrome P450cam. J Mol Biol 195, 687-700
9
Zhou, D. J., Pompon, D. and Chen, S. A. (1991) Structure-function studies of
human aromatase by site-directed mutagenesis: kinetic properties of mutants
Pro-308→Phe, Tyr-361→Phe, Tyr-361→Leu, and Phe-406→Arg. Proc Natl
Acad Sci U S A 88, 410-414
10
Koop, D. R., Laethem, C. L. and Tierney, D. J. (1989) The utility of pnitrophenol hydroxylation in P450IIE1 analysis. Drug Metab Rev 20, 541-551
11
Peter, R., Bocker, R., Beaune, P. H., Iwasaki, M., Guengerich, F. P. and
Yang, C. S. (1990) Hydroxylation of chlorzoxazone as a specific probe for
human liver cytochrome P-450IIE1. Chem Res Toxicol 3, 566-573
12
Kharasch, E. D., Thummel, K. E., Mhyre, J. and Lillibridge, J. H. (1993)
Single-dose disulfiram inhibition of chlorzoxazone metabolism: a clinical
probe for P450 2E1. Clin Pharmacol Ther 53, 643-650
13
Lucas, D., Ferrara, R., Gonzalez, E., Bodenez, P., Albores, A., Manno, M.
and Berthou, F. (1999) Chlorzoxazone, a selective probe for phenotyping
CYP2E1 in humans. Pharmacogenetics 9, 377-388
14
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J., Gonzalez, F. J. and
Miners, J. O. (1993) Validation of 4-nitrophenol as an in vitro substrate probe
for human liver CYP2E1 using cDNA expression and microsomal kinetic
techniques. Biochem Pharmacol 46, 1975-1981
43
Chapter 3
Site-Directed Mutagenesis, Expression, and Protein Purification
Introduction
To help elucidate the structural foundations for the substrate
specificities of the human 2A and 2E CYP450 subfamilies, site-directed
mutagenesis was used to exchange dissimilar active site residues between
CYP2E1, CYP2A13, and CYP2A6. Twelve differing amino acids located in the
SRS-1 and SRS-2 regions of CYP2E1 were chosen for individual
mutagenesis. The 12 desired mutations were CYP2E1 L103Q, P104A,
A105T, H107D, H109F, D111G, R112Y, W122A, P213S, W214T, L215G, and
P222S. To replace the entire SRS-1 region of CYP2E1 with that of CYP2A13,
a series of 3 sequential mutagenesis reactions were used, each altering
multiple codons. Each altered enzyme was then expressed in E. coli Topp-3
cells and purified so that the specific enzymatic activity for each CYP2E1
mutant could be determined.
Site-directed mutagenesis is a useful molecular biology technique that
can be used to introduce a desired mutation at a specific site within a DNA
sequence of interest. This technique is used to purposefully modify a gene by
substituting, inserting, or deleting, single or multiple nucleotide base pairs at a
44
desired point in the DNA sequence to determine the contributions of the
encoded amino acid(s) to protein structure and function. There is a range of
site-directed mutagenesis methods that can be used to accomplish this goal
[1-3], but many of these methods are an adaptation on a single strategy,
described below and in Figure 3.1.
In order to perform site-directed mutagenesis, it is necessary to know
the nucleotide sequence of the gene to be mutated. This is often
accomplished through DNA sequencing. To alter your nucleotide sequence, a
DNA fragment of ~20-50 base pairs in length, must be designed and
chemically synthesized. This oligonucleotide is complementary to the parental
DNA sequence at all of the base pairs except the codon encoding the amino
acid that is to be altered. The next step is to separate the two complementary
strands of the parental DNA (Figure 3.1, Step 2). This can be accomplished
by cloning the nucleic acid sequence to be mutated into a single-stranded
vector such as M13, fd, or f1 [4-8], or by separating the strands using
increased temperature. As illustrated in Step 3, the oligonucleotide is added
to the single stranded DNA and conditions altered to promote annealing to a
single parent strand (step 4). DNA polymerase can then use the
oligonucleotides as a primer to extend the DNA sequence so that a
complementary strand is formed. If the DNA sequence is replicated as shown
(Figure 3.1, step 5), two strands of double stranded DNA are formed; one
45
Figure 3.1: Basic site-directed mutagenesis strategy. Red indicates the parental
DNA sequence, black indicates the altered DNA sequence, blue indicates the
oligonucleotide, and green indicates the newly synthesized DNA.
46
resembles the original unaltered gene, and the other has incorporated the
newly designed sequence embedded within it. If the genetic sequence of
interest is generated downstream of the appropriate transcriptional and
translational elements, the resulting plasmid can be introduced into an
organism like E. coli where the mutated sequence can be expressed to yield
protein with the desired modifications. The modified protein can be used in
studies to determine the contributions of the modified amino acid(s) to protein
structure and function, but usually must first be purified from the mixture of
host proteins.
Numerous variations of the site-directed mutagenesis procedure have
been reported [9]. However, many call for a single-stranded DNA (ssDNA)
template and are often relatively lengthy, labor-intensive and have decreased
mutagenic efficiency. The QuikChange® II site-directed mutagenesis strategy
was chosen to exchange the CYP2E1 active site residues since it is a rapid
and highly efficient method that does not require the isolation of ssDNA. The
QuikChange® II method can rapidly modify the DNA sequence to exchange,
insert, or delete one or more amino acids with 80% efficiency or greater,
according to the manufacturer (Stratagene). The QuikChange® II site-directed
mutagenesis strategy makes use of a double-stranded DNA (dsDNA) vector
containing the genetic sequence of interest and two complementary synthetic
47
oligonucleotide primers that encode the desired altered nucleotide sequence
(Figure 3.2).
The method employs a three-step thermal cycling reaction to synthesize
the full plasmid nucleotide sequence while incorporating the mutation. The
first step denatures the dsDNA plasmid template by using high temperatures.
The second step lowers the temperature to anneal the synthetic mutagenic
oligonucleotides to the template DNA. The final step raises the temperature
slightly to the ideal temperature for polymerase activity, thereby extending the
annealed oligonucleotide primers.
The thermal cycling reaction is followed by a Dpn I digestion, which
selects for the newly synthesized DNA by digesting both methylated and
hemimethylated DNA. Nearly all E. coli strains methylate their DNA. Thus, the
parental DNA is susceptible to digestion, but the PCR-synthesized DNA is not
methylated and is resistant to Dpn I cleavage. The DNA is then transformed
into XL1-Blue supercompetent E. coli cells for replication and DNA nick repair.
Figure 3.2 is a simple illustration of the methods employed by the
QuikChange II strategy.
48
Figure 3.2: A schematic representation of the site-directed mutagenesis strategy
utilized by the Stratagene QuikChange II site-directed mutagenesis kit. (Stratagene)
49
The mutagenesis control provided by the QuikChange II site-directed
mutagenesis kit, is the pWhitescriptTM 4.5-kb control plasmid. This plasmid
includes the genetic sequence for β–galactosidase containing a key mutation
eliminating its activity. When the control primers are used to create a point
mutation in the plasmid, the stop codon (TAA) at amino acid 9 is converted to
the original glutamine codon (CAA). If the mutagenesis reaction was
successful and the stop codon was replaced, the resulting E. coli colony will
express
β-galactosidase
and
convert
5-bromo-4-chloro-3-indolyl-β-D-
galactopyranoside (X-Gal) into an indigo when the inducer, isopropyl-1-thio-βD-galactopyranoside IPTG, is present. If the mutagenesis reaction was
unsuccessful and the stop codon was not replaced with the glutamine codon,
the E. coli colony will not express β-galactosidase and be unable to convert XGal into an indigo. Therefore, the resulting XL1-Blue colony will appear white.
This mutagenesis control reaction is carried out in parallel to that of the
sample reactions. The ratio of blue to white colonies that grow on the control
LB-ampicillin plates provides a facile estimate of the efficiency of the control
mutagenesis reaction, which is then used to estimate the efficiency of any
parallel sets of mutagenesis reactions in the desired construct.
After transformation into E. coli cells for DNA expansion, mutant
plasmids were purified using the Wizard® Plus Minipreps DNA purification
strategy
(Promega
Corp.).
The
Wizard®
Miniprep
is
a
quick
and
50
straightforward method for isolating plasmid DNA, in which the plasmid DNA
is purified via binding to a silica resin in high salt conditions. Restriction
enzyme digestion and DNA sequencing were both used to confirm the
incorporation of the designed mutation into the genetic sequence for CYP2E1
and verify the absence of any unexpected mutations.
The CYP2E1 and subsequent mutant plasmids were then transformed
and expressed in Topp-3 E. coli cells. Following the addition of aminolevulinic
acid
(ALA) and isopropyl-1-thio-β-D-galactopyranoside (IPTG) to the
incubation mixture, E. coli cells were grown for an additional 48 hours. ALA is
an essential building block for the heme porphyrin ring found in all CYP450s.
IPTG was used to induce gene expression by preventing the lac repressor
from interfering with the transcription of the gene by the RNA polymerase.
After the E. coli cells were harvested, cells were lysed and fractionated
in preparation for column chromatography. The lysis was accomplished by
exposure to lysozyme and a hypotonic solution, and sonication. Membranebound CYP450 enzymes were solubilized from the bacterial membrane using
a detergent. Purification of the enzyme was accomplished using Fast Protein
Liquid Chromatography (FPLC). FPLC is an automated form of column
chromatography that is capable of running at moderate pressure flow rates.
51
For our purposes, we utilized two columns: the nickel-nitrilotriacetic acid (NiNTA) metal affinity and the carboxymethyl (CM) ion-exchange columns.
The Ni-NTA purification method can be divided into 5 stages (Figure
3.3). During the first stage in the procedure, the Ni-NTA resin is packed and
equilibrated with loading buffer. In the second stage, the cell lysis solution is
loaded onto the column where the non-native protein His residues (His-tag)
binds reversibly to the Ni2+. The third stage involves washing the column with
small quantities of the competing histidine side chain (8 mM imidazole) to
remove any weakly binding molecules. During the fourth stage, the imidazole
concentration is increased to elute the CYP450 enzyme. The final stage
involves regeneration of the column back to the starting conditions.
52
Figure 3.3: A schematic of Ni-NTA metal affinity column chromatography.
Stage 1
Stage 2
Equilibrate
Load
Stage 3
Wash
H
H
H
Ni2+
Ni2+
Ni
Ni2+
Ni
Ni
Ni2+
Ni2+ I
I
Ni
2+
Ni
Ni2+
H
Ni2+
2+
H
I
Ni
I
I
Ni2+
H
I
Ni2+
Ni2+
2+
Ni2+
Ni2+
Regeneration
I
Ni2+ H
Ni2+
H
2+
2+
H
H
2+
Ni
I
I
2+
Stage 5
Elution
I
H
Ni2+
Stage 4
Ni2+
Ni2+
Ni2+
I
I
H
H
H
I
I
H
H
= His-tagged protein
I
= Imidazole
= Contaminant
53
The CM ion-exchange method can also be divided into five stages
(Figure 3.4). The first stage involves establishing the column conditions for
protein binding, in terms of pH and ionic strength. During the second stage,
the sample is loaded onto the column and the positively charged protein of
interest is bound reversibly to the negatively charged carboxymethyl groups.
Substances that are not bound are washed out using the loading buffer. The
protein is eluted from the resin during the next stage by altering the conditions
to those that are unfavorable for ionic bonding. This involves increasing the
ionic strength of the buffer. Molecules elute in order of their binding
strength.The fourth and fifth stages are cleaning and returning the column to
the starting conditions.
54
Figure 3.4: A schematic of ion-exchange column chromatography.
Stage 1
Stage 2
Equilibrate
Stage 3
Load
+
+
Stage 4
Wash
Elution
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
+
Regeneration
+
+
+
Stage 5
= Positively charged protein + = Strongly charged ion
+
+
= Contaminant
55
Methods
Site-Directed Mutagenesis
The pKK233-2 expression vector (Pharmacia, Stockholm, Sweden)
including the cDNA for human CYP2E1dH and an ampicillin resistant gene
was used as the template for each mutagenesis reaction. Each of the
synthetic oligonucleotide primers was designed to be complementary to
opposite strands of the genetic sequence of interest and include the altered
nucleotide sequence required to encode the desired amino acid mutation. A
silent mutation was engineered near the altered sequence to insert or delete a
cut site for a specific restriction enzyme.
The individual mutagenic primers were designed according to the five
guidelines suggested by the manufacturer. Specifically, each oligonucleotide
was constructed with 1.) the altered genetic sequence located in the center of
the primer with 10 to 15 complementary base pairs on either side (totaling
between 25 to 45 base pairs in length), 2) a melting temperature greater than
78 oC, 3.) modest secondary structure, 4.) a GC content of greater than 40%,
and 5.) one or more G or C bases at each end of the primer. The melting
temperatures were predicted by using the equation:
56
Tm = 81.5 + 0.41(%GC)-675/N - % mismatch, where N equals the number of
bases in the oligonucleotide.
The DNA polymerase, PfuUltraTM high-fidelity (HF) DNA polymerase,
extended the oligonucleotide primers during thermal cycling using a PCR
Sprint Thermal Cycler (Thermo Scientific Inc., Waltham MA). Each
temperature cycling reaction was divided into 2 segments (see Table 3.1).
The first segment included a 30 second denaturing cycle at 95 oC. The
second segment involves a second 30 second denaturing step at 95 oC, a 1
minute cycle at 55 oC to anneal the primers, and a 1 minute/kb of plasmid
cycle at 68 oC to allow for extension of the DNA sequence. This second
segment was then repeated a total of 16 times.
Following the temperature cycling, an hour long digestion at 37 oC with
the Dpn I restriction enzyme was carried out to remove the parental and
hybrid parental/mutant DNA.
57
Table 3.1: The thermal cycling parameters used for
mutagenesis reactions.
Segment
Cycles
Temperature
Time
1
1
95 oC
30 seconds
95 oC
30 seconds
55 oC
1 minute
2
16
68 oC
1 minute/kb of
plasmid length
58
The DNA was transformed into XL1-Blue supercompetent cells by heat
shock for 45 seconds at 42 oC in BD Falcon® 2059 polypropylene roundbottom tubes (15 mL). The cells were then allowed to recover for 1 hour at 37
o
C, in 500 µL of NZY+ broth. A 250 µL aliquot of the transformation reaction
was plated and grown on Luria-Bertani (LB)-ampicillin agar plates containing
80 μg/ml 5-bromo-4-chloro-3-indolyl-β-D-galactopyranoside (X-gal) and 20
mM isopropyl-1-thio-β-D-galactopyranoside (IPTG).
The mutagenesis control described earlier (pWhitescriptTM 4.5-kb control
plasmid) was carried out in parallel to that of the sample reactions. The
number of blue to white colonies grown on the LB-ampicillin plates was used
to estimate the efficiency of the parallel sets of CYP450 mutagenesis
reactions.
Purification of Plasmid DNA
Four to six individual XL1-Blue colonies were isolated from the LBampicillin agar plates and grown overnight at 37 oC in a 5 mL LB-ampicillin
culture. The cells were pelleted by centrifugation and the supernatant
decanted. The plasmid DNA was purified using the Promega Wizard® Plus
Minipreps DNA Purification System, according to the manufacturer’s protocol.
59
Restriction Enzyme Digestions
The purified plasmids were digested using the optimal reaction
conditions recommended for the restriction enzyme whose recognition site
would be altered by the designed silent mutation. See Table 3.2 for a list of
each mutant, their restriction enzymes, and the reaction conditions. Following
the
restriction
enzyme
digestion,
the
DNA
was
analyzed
by
gel
electrophoresis (8% agarose) to determine the number and size of DNA
fragments generated from the digestion. Each gel also included a 1 kb DNA
ladder as a size reference and a negative control (uncut wild type plasmid).
The samples that contained the number and size of the DNA fragments
expected for the mutant sequence were sent to Idaho State University for full
sequencing of the CYP2E1 gene to confirm the presence of the desired
mutation and the absence of any unexpected mutations.
60
Table 3.2: The restriction enzyme reaction conditions for each CYP2E1 mutant
with the corresponding restriction enzyme.
CYP450
2E1
MUTANTS
RESTRICTION
ENZYME
L103Q
BstE II*
P104A
BstE II*
A105T
BstE II*
H107D
Bsm I**
H109F
Bsm I**
D111G
Sac II*
R112Y
Fsp I**
W122A
Sfc I**
P213S
Pst I***
W214T
Pst I***
L215G
Nco I*
P222S
Bpm I**
SRS1 rxn 1
Dra I**
SRS1 rxn 2
Bsm I**
SRS1 rxn 3
Bsm I**
REACTION
CONDITIONS
Buffer D + BSA,
1 hr at 60 oC
Buffer D + BSA,
1 hr at 60 oC
Buffer D + BSA,
1 hr at 60 oC
NEBuffer 2, 1 hr
at 65 oC
NEBuffer 2, 1 hr
at 65 oC
Buffer C + BSA,
1 hr at 37 oC
NEBuffer 4, 1 hr
at 37 oC
NEBuffer 4, 1 hr
at 37 oC
Buffer H + BSA,
1 hr at 37 oC
Buffer H + BSA,
1 hr at 37 oC
Buffer D + BSA,
1 hr at 37 oC
NEBuffer 3, 1 hr
at 37 oC
NEBuffer 4, 1 hr
at 37 oC
NEBuffer 2, 1 hr
at 65 oC
NEBuffer 2, 1 hr
at 65 oC
SIZE OF DNA FRAGMENTS (bp)
WILD TYPE
MUTANT
6038
circular
6038
circular
6038
circular
6038
linear
6038
linear
6038
circular
98, 1521,
2134, 2285
191, 678,
5169
6038
circular
6038
circular
6038
linear
1428, 2052,
2558
19, 692,
5327
6038
linear
4529, 1509
6038
linear
6038
linear
6038
linear
4529, 1509
4529, 1509
6038
linear
98, 2134
3806
191, 678,
2227, 2942
6038
linear
6038
linear
578, 5459
666, 1428,
1892, 2052
19, 92,
1913, 3414
4529, 1509
6038
linear
* Restriction enzyme and reagents supplied by Promega Corp.
** Restriction enzyme and reagents supplied by New England Biolabs Inc.
*** Restriction enzyme and reagents supplied by Fisher BioReagents.
61
Protein Expression
The pKK2E1dH wild type and mutant plasmids were each transformed
into tetracycline-resistant E. coli Topp-3 cells (Stratagene, La Jolla, CA) by
heat shock at 42 oC for 30-45 seconds. The cells were incubated in SOC
media at 37 oC, 250 rpm for 1 hour, harvested by centrifugation, and
resuspended in approximately 100 µL of the supernatant. The transformation
reactions were plated on LB-ampicillin plates and incubated overnight at 37
o
C. One colony was used to inoculate 5 mL of LB media supplemented with
135 µM ampicillin and 52 µM tetracycline. The starter culture was incubated
for 8 hrs at 37 oC and 250 rpm. 50 µL of the starter culture was used to
inoculate a 200 mL overnight culture of the LB-ampicillin-tetracycline media.
15 mL of the 200 mL overnight culture was then used to inoculate 250 mL of
Terrific Broth (TB) media, including 135 µM ampicillin, at 37 oC and 250 rpm,
and grown to an OD600 of 1.0-1.5 (approximately 2 hours). The expression of
human CYP2E1dH proteins were induced by the addition of 1 mL of 150 mM
5-aminolevulinic acid ALA and 250 mM IPTG. Human CYP2E1dH and mutant
proteins were expressed with an induction time of 48 hours at 30 oC and 190
rpm.
62
Protein Purification
The Topp-3 E. coli cells were harvested and disrupted as previously
described. After centrifugation to remove cell debris, 0.5% Na cholate was
added to the supernatant mixture and stirred on ice for 30 minutes to
solubilize the CYP450 protein, and then ultracentrifuged at 30,000 rpm for 60
min to pellet membranes. The solubilized CYP450 protein was applied to a
Ni2+-NTA Sepharose® CL-6B resin (Qiagen, Hilden, Germany), which was
subsequently washed with loading buffer. The resin was washed first with 2
column volumes of 100 mM potassium phosphate buffer (100 mM potassium
phosphate, pH 7.4, 20% glycerol, 300 mM NaCl, 0.5% Na cholate) and then
with 3 column volumes of the same buffer including 8 mM imidazole and a
reduced concentration of NaCl to 200 mM. The CYP450 protein was then
eluted with 4 column volumes of 10 mM potassium phosphate buffer with
NaCl reduced to 100 mM, and the addition of 10 mM EDTA and 250 mM
imidazole. The CYP450-containing fractions were pooled, diluted 10X with 5
mM potassium phosphate, pH 7.4, 20% glycerol, 1 mM EDTA and 0.5% Na
cholate, and loaded onto a CM-sepharose CL-6B column (GE Healthcare,
Uppsala, Sweden). The CM column was washed with 10 column volumes of
the loading buffer. Purified CYP450 protein was eluted using 4 column
volumes of the loading buffer but with potassium phosphate and NaCl
increased to 50 mM and 500 mM, respectively. The elution fractions
63
containing purified CYP450 were pooled, quantitated using a reduced CO
difference assay according to Omura and Sato [10, 11], and stored at -80oC.
Results
The mutagenic primers designed to modify the CYP2E1dH gene are
shown and each of their physical characteristics are listed in Table 3.3. Table
3.4 includes an estimate of cellular growth and other site-directed
mutagenesis results.
Restriction enzyme digestions were used to screen for the colonies
that contained the desired mutant plasmids as described in the methods. By
this method of mutant screening, we had 69% mutagenic efficiency overall.
The entire CYP2E1 gene was fully sequenced for each to confirm that the
single point mutations were present and that the plasmid contained no
unexpected mutations.
64
65
CTCGGGCAGAGGTGACCTCGCCGCGTTCCATGC
CTCGGGCAGAGGTGACCTCCCCACCTTCCATGC G C A C
GGCGACCTCCCCGCATTCGATGCGCACAGGGAC
CCTCCCCGAGTTCCATGCATTCAGGGACAGGGGAATC
GCGTTCCATGCGCACCGCGGCAGGGGAATCATTTTTAATAA
TGG
CCGCGTTCCATGCCCACAGGGACTATGGAATCATTTTTAATA
ATGGACC
GGAATCATTTTTAATAATGGACCTACAGCGAAGGACATCCG
GCGG
CCTACTCAGCACTTCCTGGCTGCAGCTTTACAATAATTTTC C
P104A
A105T
H107D
H109F
D111G
CCTACTCAGCACTCCATGGGGCCAGCTTTACATTATTTTTC C
GCTCCAGCTTTACAATAATTTCTCCAGCTTTCTACACTACTT
GCC
GCAGAGGCGACCTCCCCGCGTTCCATCTCTTTAAAGGCTAT
GGAATCATTTTTAATAATGG A C C
CGAGTTCTCGGGCAGAGGCGAACAAGCTGCATTCCATCTCT
TTAAAGGCTATGG
GGCGAACAAGCTACCTTCGATTGGCTCTTTAAAGGCTATG G
L215G
P222S
SRS1 rxn
1
SRS1 rxn
2
SRS1 rxn
3
CTACTCAGCACTCCCACGCTGCAGCTTTACAATAATTTTCC
W214T
P213S
W122A
R112Y
TTCTCGGGCAGAGGTGACCAGCCCGCGTTCCATGC C
Sequence (5! to 3!)
L103Q
Mutation
Length
64
54
41
Additional BsmI
Removal BsmI
45
42
42
42
45
49
44
37
33
37
33
36
(base pairs)
Additional DraI
Additional GsuI
Additional NcoI
Additional PstI
Additional PstI
Additional SfcI
Removal FspI
Additional SacII
Additional BsmI
Additional BsmI
Additional BstEI I
Additional BstEI I
Additional BstEI I
Restriction
site altered
85.0
79.3
82.0
79.4
78.1
77.8
79.2
79.0
78.8
80.6
80.7
83.6
82.9
83.6
82.1
Tm
(oC)
48.78
51.85
46.88
42.22
47.62
47.62
45.24
46.67
49.94
52.27
62.16
69.70
67.57
69.70
66.67
(%)
GC
Moderate
Strong
Strong
Moderate
Weak
Strong
Strong
Weak
Moderate
Moderate
Very Strong
Strong
Strong
Strong
Very Strong
Sec. S t r .
No
No
No
No
No
No
No
No
No
No
No
No
No
No
No
Primer
Dime r
Table 3.3: Sequences for one of the two oligonucleotides used in the creation of each of the CYP2E1 mutations. The second
oligonucleotide was complementary to the oligonucleotide sequence shown. Each col ored residue denotes modifications of the wild type
sequence. Blue denotes the position of the desired mutation. Red denotes an alteration that modifies a restriction site.
Table 3.4: Site-directed mutagenesis results. Scoring for number of colonies per LB
plate: * 1-10 colonies, *** = 30-50 colonies, ***** = plate fully covered. DNA
sequencing results that confirmed no unexpected mutations are indicated by Okay.
N/A represents mutagenesis reactions that were not sequenced.
CYP450
Mutant
SDM
(50ng
and
20ng)
Dpn1
Digestion
Transformation
into XL-1 Blue
Super
Competent
Cells
(50ng/20ng)
Control
(Ratio of
White:Blue
colonies)
Mutagenic
Efficiency
(# mutant
colonies: #
colonies
screened)
Sequencing
Results
L103Q
√
√
***/**
1:10
7:9
Okay
P104A
√
√
***/***
1:6
3:3
Okay
A105T
√
√
****/***
1:6
4:6
Okay
H107D
√
√
***/***
1:6
3:4
Okay
H109F
√
√
**/*
1:6
2:5
Okay
D111G
√
√
**/*
1:6
4:4
Okay
R112Y
√
√
***/***
1:8
5:16
Okay
W122A
√
√
***/***
1:8
2:3
W122A +
D111G
P213S
√
√
***/**
1:4
3:4
Okay
W214T
√
√
***/***
1:4
1:4
Okay
L215G
√
√
***/***
1:4
3:4
Okay
P222S
√
√
***/**
1:4
3:4
Okay
W122A
√
√
**/**
1:4
4:4
Okay
SRS1
rxn1†
√
√
***/**
1:5
5:8
N/A
SRS1
rxn2††
√
√
****/**
1:7
4:7
Okay
SRS1
rxn3
√
√
****/***
1:15
3:4
Okay
†SDM carried out with 50 ng and 30 ng of plasmid DNA.
††SDM carried out with 50nm and 25 nmg of plasmid DNA.
66
Through this approach we were able to successfully obtain each of the
designed point mutations. The multiple amino acid mutations required to
replace the entire SRS-1 region of CYP2E1 were designed in a series of three
mutagenesis reactions replacing three to four amino acids at a time. The first
reaction utilized the CYP2E1 R112Y mutant as the template DNA and
replaced three amino acid residues, and at the same time introduced a new
restriction enzyme cut site for Dra I. The three mutations were H109F, R110K,
and D111G. The second reaction created three additional amino acid
substitutions and introduced a new cut site for the restriction enzyme, Bsm I.
The three amino acids to be exchanged in this second round of mutagenesis
were D102E, L103Q, and P104A.
The third reaction was designed to
substitute additional two amino acids, insert one amino acid present in the
CYP2A enzymes and not the CYP2E1 enzyme, and remove the Bsm I
restriction enzyme cut site introduced in the previous reaction. The two amino
acids exchanged were, A105T and H107D. A tryptophan was inserted
between amino acids 107 and 108. The following schematic represents the
changes made to the nucleic acid base pairs during each reaction to make
the desired amino acid changes. The base pairs in bold represent the base
pairs that were altered from the previous sequence.
67
Beginning A.A. Seq.:
D102 L103 P104 A105 F106 H107 A108 H109 R110 D111 Y112
Template: 5′… GAC CTC CCC GCG TTC CAT GCC CAC AGG GAC TAT… 3′
Reaction 1: 5′… GAC CTC CCC GCG TTC CAT CTC TTT AAA GGC TAT… 3′
Reaction 2: 5′… GAA CAA GCT GCA TTC CAT CTC TTT AAA GGC TAT… 3′
Reaction 3: 5′… GAA CAA GCT ACC TTC GAT CTC TTT AAA GGC TAT… 3′
TGG (insertion of W108)
Final A.A. Sequence:
E102 Q103 A104 T105 F106 D107 L109 F110 K111 G112 Y113
In addition, during the sequencing of one of the colonies, we
discovered an unexpected double mutation to D111G/W122A. This double
mutation was expressed, purified, and further analyzed by metabolism assays
along with the intended mutants.
After the transformation and expression of each of the CYP450
mutants, the E. coli cells were harvested and the fourteen CYP2E1 mutant
proteins purified using the two column purification previously described.
Purified protein was characterized and quantitated by a reduced CO
difference spectrum. Table 3.5 shows the results of this characterization for
each purified CYP2E1 protein. An absolute spectrum is read to estimate the
total CYP450 present by measuring the λmax at ~418 nm (absorbance for a
non-reduced heme with water bound in the sixth coordinate position). The
purity is determined by comparison of the absorbance band at ~418 to ~280
nm. Because aromatic amino acids absorb light at ~280 nm, the absorption at
this wavelength can be used as an estimate of purity by comparison to the
68
Table 3.5: Characterization of purified CYP2E1 proteins by UV/Vis spectroscopy and
CO difference spectra, to determine the yield, purity, and active/inactive state of each
CYP450 2E1 mutant.
λ max of
2E1 Protein
Absolute
Spectrum
(nm)
Estimated
Purity
Yield
CO Diff
(A420/280)
(nmol
CYP450/
1.25 L E. coli)
(% P450)
2E1
417
0.84
249
90%
L103Q
425
N/A
53
0%
425
1.09
195
10%
H109F
423
0.80
56
0%
R112Y
425
1.20
509
10%
W122A
424
1.20
21
0%
L215G
422
0.77
63
5%
D111G/W122A
422
1.00
55
0%
SRS-1
420
0.13
13
0%
399
N/A
93
0%
P104A
424
1.20
598
100%
A105T
424
1.04
355
80%
H107D
424
1.11
85
100%
D111G
424
1.18
399
100%
P213S
423
1.14
326
100%
W214T
422
1.15
183
95%
P222S
422
1.03
482
100%
Expression #1
L103Q
Expression #2
Expression #1
SRS-1
Expression #2
69
418 nm absorption band. The total active CYP450 present is quanitated by a
CO difference spectrum.
Seven of the mutated 2E1 proteins were isolated in the catalytically
inactive state, characterized by a λmax in CO difference spectrum at 420 nm:
L103Q, H109F, R112Y, W122A, L215G, D111G/W122A, and SRS-1. In
contrast, catically active P450 proteins have a λmax at 450 nm. The seven
CYP2E1 mutant proteins isolated in their active state were: P104A, A105T,
H107D, D111G, P213S, W214T, and P222S. The loss of activity in half of the
mutant proteins could be due to the original amino acid’s contributions to
overall protein stabilization.
Of note is that the SRS-1 mutant was unable to be purified (A41/280 =
0.13). This implies that the SRS-1 region in the 2A subfamily could have a
different conformation(s) than that of the 2E subfamily. A significant difference
found between the two SRS-1 regions in these subfamilies is at amino acid
residue 108. The amino acid residue 108 in the 2A subfamily is a conserved
tryptophan. However in the 2E subfamily, this amino acid residue is deleted.
This extra residue could be a key player in the destabilization of the CYP2E1
protein when the entire 2A SRS-1 region has been inserted.
70
Conclusion
The QuikChange mutagenesis afforded a quick and easy method for
performing the desired changes to the CYP2E1 DNA sequence. All 12 of the
single amino acid mutations were easily obtained as well as one unexpected
mutation, D111G/W122A. The multiple amino acid mutation to replace the
entire SRS-1 region of CYP2E1 was easily accomplished in a series of three
sequential reactions replacing 3-4 amino acids at a time. The overall
mutagenic
efficiency
estimated
by
restriction
enzyme
digest
was
approximately 69%.
Purification of these fourteen mutated proteins by metal affinity and ion
exchange chromatography yielded seven CYP2E1 mutant proteins in their
active form. These were mutant proteins P104A, A105T, H107D, D111G,
P213S, W214T, and P222S. However, seven of the mutant proteins were
purified in the inactive state. These were the CYP2E1 mutant proteins L103Q,
H109F, R112Y, W122A, L215G, SRS-1, and D111G/W122A. The most likely
reason that these CYP2E1 mutant proteins were in the inactive state is
disruption of the altered amino acid’s role in overall protein stabilization.
71
References
1
Deng, W. P. and Nickoloff, J. A. (1992) Site-directed mutagenesis of virtually
any plasmid by eliminating a unique site. Anal Biochem 200, 81-88
2
Horton, R. M., Hunt, H. D., Ho, S. N., Pullen, J. K. and Pease, L. R. (1989)
Engineering hybrid genes without the use of restriction enzymes: gene
splicing by overlap extension. Gene 77, 61-68
3
Kunkel, T. A. (1985) Rapid and efficient site-specific mutagenesis without
phenotypic selection. Proc Natl Acad Sci U S A 82, 488-492
4
Greenstein, D. and Besmond, C. (2001) Preparing and using M13-derived
vectors. Curr Protoc Mol Biol Chapter 1, Unit1 15
5
Yu, Q. (1996) Cloning into M13 bacteriophage vectors. Methods Mol Biol 58,
343-348
6
Dotto, G. P., Enea, V. and Zinder, N. D. (1981) Functional analysis of
bacteriophage f1 intergenic region. Virology 114, 463-473
7
Dotto, G. P. and Horiuchi, K. (1981) Replication of a plasmid containing two
origins of bacteriophage. J Mol Biol 153, 169-176
8
Herrmann, R., Neugebauer, K., Pirkl, E., Zentgraf, H. and Schaller, H. (1980)
Conversion of bacteriophage fd into an efficient single-stranded DNA vector
system. Mol Gen Genet 177, 231-242
9
Trower, M. K. (1996) Methods in Molecular Biology. Humana Press Inc.,
Totowa, New Jersey
10
Omura, T. and Sato, R. (1964) The Carbon Monoxide-Binding Pigment of
Liver Microsomes. I. Evidence for Its Hemoprotein Nature. J Biol Chem 239,
2370-2378
11
Omura, T. and Sato, R. (1962) A new cytochrome in liver microsomes. J Biol
Chem 237, 1375-1376
72
Chapter 4
Characterization of CYP2E and 2A Proteins Using Chlorzoxazone and
p-Nitrophenol Hydroxylation Assays
Introduction
It is well known that together the 2A and 2E CYP450 subfamilies have
both identical and diverse metabolic substrates [1-3]. To identify the amino
acid residues that distinguish the metabolic activities of these two CYP450
subfamilies, likely active site amino acids were exchanged between CYP2E1
and CYP450 2A enzymes as described in the previous chapter. To determine
if any of these residues are essential for CYP2E1 substrate metabolism, we
selected substrates that were reportedly selective for the 2E enzymes. We
hypothesized that mutant proteins with significant deviation from the activity of
the wild type CYP450 2E1 enzyme would indicate the mutated amino acid
makes an important interaction with the substrate that distinguishes the
metabolic capabilities of the two subfamilies.
p-Nitrophenol (pNP) and chlorzoxazone (CZN) were selected as the
CYP2E1-selective substrates based on previous studies in the literature [4-8].
pNP and CZN are both well known, low Km substrates for CYP2E1.
Traditionally, these two compounds have been used to indicate the presence
73
and relative activity of CYP2E1 in microsomal samples containing multiple
CYP450 enzymes.
Identifying a selective substrate for individual CYP450 enzymes is
important for carrying out a correlation study. Correlation studies utilizing
selective
substrates
are
commonly conducted
using
human
liver
microsomes to elucidate the CYP450 enzymes responsible for the in vitro
metabolism of a chosen substrate [9-13]. If the presence of a compound
under investigation represses the metabolism of a CYP450-selective
substrate, the conclusion is drawn that the CYP450 enzyme that
metabolizes the selective substrate also binds the compound being
evaluated. If there are multiple CYP450 enzymes
capable of
metabolizing a supposedly “selective substrate,” this conclusion is not
valid and determinations must be made with isolated or purified CYP450
to establish metabolism.
In order to metabolize any substrate, each purified CYP450 enzyme
must be reconstituted with associated electron delivery proteins [14]. In our
studies, a reconstitution mixture of 1:6:2 for CYP450:NADPH-P450
oxidoreductase:cytochrome b5 was found to support maximal metabolism.
Nicotinamide adenine dinucleotide phosphate (NADPH) is added to each
reaction as the electron source in sufficient quantities to initiate and support
metabolism, until the reaction is halted by the addition of a precipitant. The
74
metabolite of each reaction is quantified and the rates of substrate
metabolism determined for each protein.
Besides its reported selectivity for CYP2E1, another advantage of
utilizing the pNP hydroxylation assay to measure CYP2E1 activity is the ease
with
which
the
pNP
metabolite
can
be
measured.
A
simple
spectrophotometric analysis is the most commonly used method to determine
the amount of pNP metabolite produced directly in the reaction mixture [4, 1517], although the metabolite can also be separated from the mixture by highpressure liquid chromatography (HPLC) prior to UV detection [18, 19].
pNP is a phenolic compound with a pKa of 7.08. A solution of the
phenolic substrate is colorless, but the phenolate salt is bright yellow with an
absorption maximum at approximately 405 nm. This colorimetric property of
pNP contributes to its use as a pH indicator. CYP2E1 hydroxylates pNP at the
2 position to generate 4-nitrocatechol (4NC), as shown in Figure 4.1. 4NC
also posses a similar colormetric property to pNP. 4NC has a pKa of 7.15,
and when present as a phenolate salt, turns a bright pinkish purple with an
absorption maximum at 510 nm.
Because of these spectral properties,
metabolism of pNP into 4NC is easily monitored using a spectrophotometer
[4]. After the reaction is terminated, a small amount of NaOH can be added to
the reaction supernatant to alter the pH of the solution so that the 4NC is
primarily found as the phenolate salt. Using Beer’s Law and a standard curve,
75
Figure 4.1: The hydroxylation of p-nitrophenol catalyzed
by CYP450 2E1 and colorimetric detection of substrate
and metabolites.
p-Nitrophenol
4-Nitrocatechol
OH
OH
OH
CYP450
+ 2e- + O2
NO2
λmax = 250 nm
+ NaOH
O-Na+
NO2
λmax = 250 nm
+ NaOH
O-Na+
O-Na+
NO2
λmax = 405 nm
NO2
λmax = 510 nm
76
the metabolite is easily quantitated and the enzymatic activity can be
determined. Alternatively, after metabolite separation from the reaction
mixture by HPLC, the 4NC metabolite can be detected directly by its λmax at
250 nm.
Chlorzoxazone is a drug that is occasionally prescribed as a muscle
relaxant to prevent muscle spasms and the ensuing pain or discomfort. The
National Cancer Institute defines chlorzoxazone as: “Highly selective for
CYP2E1, CZN may be used as a selective probe for phenotyping CYP2E1 in
humans; the ratio of 6OH-CZN to CZN plasma concentration obtained 2-4
hours after oral administration of CZN may be used as a phenotypic measure
of CYP2E1 enzymatic activity. [20]” In the human body, this drug is
hydroxylated by CYP2E1 into 6-hydroxychlorzoxazone (6OH-CZN), which
subsequently undergoes phase II glucuronidation and is excreted renally [20].
Figure 4.2 is an illustration of the hydroxylation of CZN by CYP2E1. The
metabolism of CZN is monitored by detection of the metabolite at a
wavelength of 287 nm following separation of reaction components by highpressure liquid chromatography (HPLC).
77
Figure 4.2: The 6-hydroxylation of chlorzoxazone catalyzed by CYP450 2E1.
Chlorzoxazone
Cl
6OH-Chlorzoxazone
H
N
Cl
O
O
λmax = 287 nm
H
N
CYP450
O
+ 2e- + O2
HO
O
λmax = 287 nm
78
HPLC is a type of column chromatography that can be used to isolate
a compound of interest by capitalizing on an assortment of interactions
between the substance being analyzed and the chromatography column. For
our purposes, a reversed-phase C18 chromatography column was used to
isolate both pNP and CZN metabolites. Reversed phase chromatography
consists of two phases: a nonpolar stationary phase and a polar mobile
phase. A typical stationary phase consists of silica covalently bonded to a
straight alkyl group such as, C18H37 or C8H17. Because of the hydrophobic
nature of the chromatography column, the retention time is extended for
molecules that are nonpolar, and reduced for more polar molecules. The
retention time of a given molecule can be increased or decreased by
changing the polarity of the mobile phase.
The introduction of a more
hydrophilic solvent will increase the retention time of a given molecule, and
the addition of a more hydrophobic solvent will decrease the retention time.
Thus, HPLC separation coupled with UV detection is a sensitive method for
the isolation, detection, and quantification of the CZN and pNP metabolites,
and determining the enzymatic activity of CYP450 enzymes that produce
these metabolites.
79
Methods
p-Nitrophenol Metabolism Assay
p-Nitrophenol 2-hydroxylation by CYP450 enzymes was determined
initially using the colorimetric properties of the phenolic salt of pNP directly in
the reaction mixture, but later was determined using HPLC separation and UV
detection because of increased reducibility. Before each assay, a reduced CO
difference spectrum was obtained in the metabolism assay buffer (100 mM
potassium phosphate, pH 6.8), for each of the thawed, purified CYP450
enzymes to quantitate the amount of active protein. To determine activities,
the incubation mixtures consisted of 30 pmol CYP450 in a reconstituted
mixture (1:6:2, CYP450:NADPH-P450 oxidoreductase:cytochrome b5), 100
mM potassium phosphate (pH 6.8), 150 µM p-nitrophenol, 1 mM ascorbic
acid, and 1 mM NADPH, in a final volume of 1 mL. After the addition of
NAPDH to initiate catalysis, the reactions were then incubated at 37 oC for 10
minutes. Each reaction was terminated by the addition of 300 µL of 20%
tricholoacetic acid to precipitate protein. Subsequent incubation on ice for 10
minutes followed by centrifugation at 3000 rpm for 15 minutes clarified the
solution. The metabolic activities were calculated from duplicate trials, each
consisting reactions run in triplicate.
80
For the colorimetric detection of pNP and 4NC in the reaction mixture,
1 mL of the supernatant, plus 100 µL of 10 M NaOH were added to a
disposable cuvette, and incubated at room temperature for 1 minute. The
absorbances values were read at a wavelength of 510 nm using an UV2501PC UV-VIS Spectrophotometer (Shimadzu Scientific Instruments, Inc.,
Kyoto, Japan).
For HPLC analysis, the LC-10A VP Prominence HPLC system
(Shimadzu Scientific Instruments, Inc., Kyoto, Japan) was used. Separation
was accomplished using a Luna 5 µ C18 (2) 100 A column (150 x 4.60 mm)
operated at 37 oC. Absorbance was monitored at a wavelength of 250 nm.
The isocratic mobile phase, delivered at a flow rate of 1.5 mL/min, was
comprised of acetonitrile-glacial acetic acid-water (22:1:77) containing 30 mM
triethylamine; the pH was adjusted to 3.0 using phosphoric acid. Fifteen µL of
the reaction supernatant was injected into the HPLC system. Standard curves
for both the spectrophotometric and HPLC analysis were generated using
seven 4-nitrocatechol standards with a concentration range of 0.5-7.5 µM.
To determine kinetic parameters, 10 reactions were run with varying
concentrations of p-nitrophenol: 0, 10, 20, 35, 50, 75, 100, 150, 250, and 350
µM. The kinetic parameters were calculated from duplicates trials, each
consisting of reactions done in triplicate.
81
Chlorzoxazone Metabolism Assay
Chlorzoxazone 6-hydroxylation activities by CYP450 enzymes were
determined by HPLC separation and UV detection. Before each assay, a
reduced CO difference spectrum was obtained in the metabolism assay buffer
(100 mM potassium phosphate, pH 7.4), for each of the thawed, purified
CYP450 mutants to quantitate the amount of active protein. To determine
activities, the incubation mixtures consisted of 30 pmol CYP450 in a
reconstituted
mixture
(1:6:2,
CYP450:NADPH-P450
oxidoreductase:
cytochrome b5), 100 mM potassium phosphate (pH 7.4), 300 µM
chlorzoxazone, 200 U superoxide dismutase, 200 U catalase, and 1 mM
NADPH, with a final volume of 500 µL. After the addition of NADPH to initiate
catalysis, the reactions were allowed to proceed at 37 oC for 10 minutes. Each
reaction was terminated by the addition of 25 µL 60% perchloric acid to
precipitate the protein. The reaction was briefly chilled on ice, and followed by
centrifugation at 5000 rpm for 10 minutes to clarify the solution. The metabolic
activities were calculated from duplicates trials, each consisting of reactions
done in triplicate.
For the HPLC analysis, the LC-10A VP Prominence HPLC system
(Shimadzu, Scientific Instruments Inc., Kyoto, Japan) was used. The samples
were run on a Luna 5 µ C18 (2) 100 A column (150 x 4.60 mm) operating at 35
o
C. The absorbance was monitored at a wavelength of 287 nm. The isocratic
82
mobile phase, delivered at a flow-rate of 1.0 mL/min, was comprised of
acetonitrile-0.05% phosphoric acid in water (20:80). Fifteen µL of the
supernatant was injected into the HPLC system. Standard curves for analysis
were prepared using seven 6-hydroxychlorzoxazone standards with a
concentration range of 0.5-10 µM.
To determine the kinetic parameters, reactions with chlorzoxazone
concentrations of 0, 25, 50, 75, 100, 150, 200, 300, and 500 µM were used.
The kinetic parameters were calculated from duplicates trials, each consisting
of reactions done in triplicate.
Results
We hypothesized that only a small number of amino acid residues are
responsible for particular substrate-enzyme interactions that differentiate the
metabolic abilities of the CYP450 2A and 2E subfamilies. To identify those
key amino acid residues, the 2-hydroxylation of p-nitrophenol and 6hydroxylation of chlorzoxazone (both marker substrates for CYP2E1 [4-8])
were determined for the human wild type CYP2E and 2A enzymes (CYP2E1,
CYP2A13, and CYP2A6) and for a set of CYP2E1 mutant proteins.
Of the fourteen CYP2E1 mutant proteins discussed in Chapter 3,
seven were isolated in their active form: P104A, A105T, H107D, D111G,
83
P213S, W214T, and P222S. These seven mutants were subsequently
examined for their ability to metabolize pNP and CZN.
p-Nitrophenol Metabolism Assay
The 2-hydroxylation of p-nitrophenol was initially determined for each
enzyme at 150 µM pNP using the colormetric assay. This concentration was
chosen because several other labs have reported a loss in metabolic activity
at concentrations higher than 200 µM pNP [15, 21, 22]. The metabolic
activities of CYP2E1, 2A13, and 2A6, were 8.36, 21.81, and 22.39
nmol/min/nmol, respectively (Figure 4.4). Unexpectedly, the CYP450 2A
subfamily enzymes had 2-2.5-fold higher pNP metabolism than CYP2E1, for
which pNP is reportedly a selective substrate. None of the seven CYP2E1
mutant proteins showed significant changes in pNP activity compared to the
wild type CYP2E1, varying only 89-116% relative to the parent wild type
enzyme activity (Figure 4.3). Although the pNP activities of the CYP2E1 and
2A enzymes are reversed from those expected, the significant difference in
pNP metabolism would still allow identification of residues that are important
in the differential metabolism of pNP. However, since none of the chosen
amino acids alone significantly alter activity, the results indicate that these
residues may not play a significant role in any enzyme-substrate interactions
distinguishing CYP2E1 from the 2A subfamily.
84
Figure 4.3: Comparison of CYP450 pNP activities (nmol/min/nmol) at 150 µM pNP.
2E1-1 and 2E1-2 indicate two different batches of purified CYP2E1 wild type
enzyme.
85
A comparison of the visible colorimetric and HPLC-based 4NC
detection methods revealed similar results for activity for each wild type
CYP450 (Table 4.1), respectively. Table 4.1 is a comparison of pNP activity
determined at 150 µM from a single run of triplicate reactions for each wild
type CYP450 enzyme. Figure 4.4 is a comparison of the p-nitrophenol kinetics
for CYP2E1 determined by both colorimetric and HPLC UV methods. Each
method was run twice and each point was run in triplicate. The colorimetric
detection had approximately a 2-fold difference in KM and kcat values between
runs. In contrast, the HPLC UV detection is much more consistent and has
similar kcat and Km values. Because the HPLC method yielded a decrease in
variability between triplicate samples and day-to-day runs, this method was
used to determine the kinetic parameters of pNP metabolism. pNP kinetics
were only determined for the wild type enzymes due to the small differences
in activity between that of the wild type CYP2E1 and mutant enzymes in the
single pNP concentration assays.
86
Table 4.1: Comparison of pNP activity (nmol/min/nmol) determined by
visible colorimeteric and HPLC UV detection methods. Activity was
determined at 150 µM p-nitrophenol.
pNP Assay
2E1
2A13
2A6
Visible
Colorimeteric
10.64 ± 0.17
21.64 ± 2.92
22.39 ± 1.15
10.15 ± 1.24
16.84 ± 1.55
23.28 ± 2.36
(nmol/min/nmol)
HPLC
UV
(nmol/min/nmol)
Figure 4.4: Comparison of Michaelis-Menten kinetics determined for
CYP2E1 by both colorimetric and HPLC UV detection methods.
10.0
7.5
5.0
2.5
0.0
0
KM
44.9 ± 19.7 µM
kcat
13.0 ± 2.6 µM-1min-1
25
50
75
100
pNP Concentration
pNP
Concentration (µM)
(µM)
Activity (nmol/min/nmol)
HPLC/UV Assay
125
7.5
5.0
2.5
0.0
0
25
50
KM
23.5 ± 8.5 µM
kcat
-1
75
6.9 ± 0.8 µM min
100
pNP Concentration (µM)
pNP Concentration (µM)
12
10
8
6
4
KM
98.2 ± 16.2 µM
2
kcat
17.3 ± 1.5 µM-1min-1
0
0
25
50
75
100 125 150 175
pNP Concentration
Concentration (µM)
(µM)
pNP
125
-1
Activity (nmol/min/nmol)
Activity (nmol/min/nmol)
Activity (nmol/min/nmol)
Day 2
Day 1
Colorimetric Assay
12
10
8
6
4
KM
146.9 ± 36.1 µM
2
kcat
16.8 ± 2.1 µM-1min-1
0
0
50
100
150
200
250
300
pNP Concentration (µM)
pNP Concentration (µM)
87
Each wild type enzyme exhibited simple Michaelis-Menten kinetics
(Figure 4.5). The Km values were similar for both CYP2E1 and CYP2A13
(75.8 and 62.7 µM), while CYP2A6 had a much higher Km of 135.8 µM.
However, the kcat for both CYP2A13 and CYP2A6 (30.3 and 52.6
nmol/min/nmol) were approximately 2 and 3-fold higher than that of CYP2E1
(15.9 nmol/min/nmol). When the catalytic efficiency of all three enzymes are
compared, CYP2A13 and CYP2A6 were both approximately twice as efficient
at catalysis of pNP than CYP2E1 (Table 4.2).
In contrast to previous literature reports for CYP2A6 [4, 15, 17, 23],
both CYP2A6 and CYP2A13 demonstrated pNP activity greater than that of
CYP2E1. Additionally, other labs have reported the appearance of inhibition
at higher concentrations of pNP (>200 µM). This was not observed in our
kinetic assays, which used concentrations of pNP up to 350 µM. This
difference could be due to one or more of the several features that differ
between our assay and the reported results (e.g., purified versus microsomal
protein, or full length versus truncated enzymes).
88
Table 4.2: The pNP kinetic parameters determined for CYP2E1,
CYP2A6, and CYP2A13.
Km
kcat
kcat/Km
(µM)
(min )
(µM min )
2E1
75.8 ± 4.4
15.9 ± 0.4
0.21
2A6
135.8 ± 13.7
52.6 ± 2.4
0.39
2A13
62.7 ± 7.4
30.3 ± 1.3
0.48
CYP450
-1
-1
-1
Figure 4.5: An overlay of the enzyme kinetics for pNP metabolism
by CYP2E1, CYP2A6, and CYP2A13.
40
30
20
10
0
0
100
200
300
400
pNP Concentration (µM)
CYP450 2E1
CYP450 2A13
CYP450 2A6
89
In conclusion, each of the purified CYP2E1 mutant proteins had little
difference in pNP activity from that of the wild type CYP2E1 protein. This may
suggest that none of the amino acids that were exchanged play a key role in
differentiating the p-nitrophenol activity of the CYP450 2E and 2A subfamilies.
On the other hand, the CYP2A enzymes CYP2A6 and CYP2A13 showed a
much higher activity for p-nitrophenol than previously reported [23]. In
particular, the increase in kcat (and decrease in Km of CYP2A13) means that
pNP metabolism cannot be reliably used as an indicator of CYP2E1 activity in
samples containing CYP450 2A enzymes.
Chlorzoxzaone Metabolism Assay
The 6-hydroxylation of chlorzoxazone was also used to characterize
activity of these wild type and mutant enzymes. Initially, the activity of each of
the enzymes was determined at a single concentration of 300 µM CZN based
on a rough kinetic assay used to approximate the kcat. At this concentration,
the activities of the wild type CYP2E1 and CYP2A6 enzymes are very similar
(Figure 4.6). However, CYP2A13 has a CZN activity that is approximately 3fold higher than that of the other two enzymes. Therefore, significant
increases in metabolic activity of any of the CYP2E1 mutants could imply that
the CYP2E1 active site has become more CYP2A13-like only and not more
like the CYP450 2A subfamily in general.
90
Figure 4.6: Comparison of CZN activities (nmol/min/nmol) of the CYP2E1 mutant
enzymes and wild type CYP450s. 2E1-1 and 2E1-2 indicate two different batches of
purified CYP2E1 enzyme. Activities were determined at 300 µM chlorzoxazone.
91
However, like the pNP results, the CYP450 mutant enzymes P104A,
A105T, H017D, D111G, and P213S all exhibited activities similar to that of the
CYP2E1 wild type enzyme (Figure 4.6). The CYP2E1 mutant enzymes
W214T and P222S demonstrated a small decrease to 60% of the wild type
CZN activity.
When CZN kinetics were performed for the three wild type enzymes,
each exhibited simple Michaelis-Menten kinetics (Figure 4.7). The Km values
were similar for both CYP2E1 and CYP2A6 (105.5 and 100.7 µM), while
CYP2A13 had a lower Km of 64.8 µM. The kcat for both CYP2E1 and CYP2A6
were also very similar (5.1 and 4.7 nmol/min/nmol), and were approximately
4-fold lower than that of CYP2A13 (22.7 nmol/min/nmol). When comparing
the enzymatic efficiency between each wild type CYP450 (Table 4.3),
CYP2A13 was approximately ten times as efficient at catalysis of CZN than
CYP2E1 and CYP2A6 (2.85 versus 20.69 and 21.24).
92
Table 4.3: CZN kinetic parameters determined for CYP2E1,
CYP2A6, and CYP2A13.
Km
kcat
kcat/Km
(µM)
(min )
(µM min )
2E1
105.5 ± 22.9
5.10 ± 0.4
0.05
2A6
100.7 ± 15.1
4.74 ± 0.3
0.05
2A13
64.8 ± 10.4
22.74 ± 1.1
0.35
CYP450
-1
-1
-1
Figure 4.7: An overlay of the enzyme kinetic parameters of CZN
CYP450
Metabolism
metabolism by CYP2E1,
CYP2A6,CZN
and CYP2A13.
20
15
10
5
0
0
100
200
300
400
500
CZN Concentration (µM)
2E1
2A6
2A13
93
In conclusion, each of the purified CYP2E1 mutants had a negligible
difference in activity relative to that of the wild type CYP2E1 protein. This
indicates that none of the amino acids that were exchanged are likely to play
a key role in differentiating the chlorzoxazone activity of CYP2E1 and
CYP2A13. Based on the higher catalytic efficiency of CYP2A13 for
chlorzoxazone and the identical catalytic efficiency of CYP2A6 to CYP2E1
means that chlorzoxazone cannot be used as a marker substrate for CYP2E1
in microsomal samples or in whole organism experiments where the CYP450
2A enzymes are also present.
Conclusion
CYP2E1, CYP2A13, and CYP2A6 each displayed differing metabolism
of both p-nitrophenol and chlorzoxazone. In particular, both the 2A enzymes
displayed a remarkable ability to metabolize p-nitrophenol over that of
CYP2E1 enzyme. In addition, the 6-hydroxylation of chlorzoxazone was
predominately metabolized by CYP2A13, while both CYP2A6 and CYP2E1
almost identically contribute to the metabolism of chlorzoxazone. As
mentioned previously, if there are multiple CYP450 enzymes capable of
metabolizing a given “selective substrate,” a proper correlation cannot
be made as to the CYP450 enzymes involved in the in vitro metabolism
of a chosen substrate. These results are important because both substrates
94
have historically been used as markers of CYP2E1 activity in mixed CYP
samples. These present results invalidate p-nitrophenol and chlorzoxazone
as selective substrates for CYP2E1.
None of the mutations in the CYP2E1 protein had significant effects on
pNP and CZN activity compared to that of the wild type CYP2E1 protein
(Table 4.4). This indicates that none of the amino acids that were exchanged
are key in differentiating the p-nitrophenol and chlorzoxazone activity of the
CYP450 2E and 2A subfamilies. Thus, other residues must be responsible for
the differences in metabolism.
95
Table 4.4: Comparison between the activities
(nmol/min/nmol) of each CYP2E1 mutant enzyme
for p-nitrophenol and chlorzoxazone.
CYP450
pNP Activity
CZN Activity
(nmol/min/nmol)
(nmol/min/nmol)
2E1-1
10.20 ± 2.63
5.40 ± 0.27
2E1-2
11.29 ± 0.92
5.56 ± 0.63
2A6
26.75 ± 2.06
5.20 ± 0.71
2A13
22.14 ± 1.40
18.30 ± 1.13
P104A
11.42 ± 0.77
5.35 ± 0.91
A105T
9.51 ± 1.31
4.12 ± 1.37
H107D
12.46 ± 0.72
7.14 ± 0.99
D111G
11.80 ± 0.14
4.80 ± 1.56
P213S
11.30 ± 2.13
5.05 ± 0.24
W214T
10.49 ± 0.83
3.27 ± 0.03
P222S
11.57 ± 0.71
3.13 ± 0.38
96
References
1
Lewis, D. F., Dickins, M., Lake, B. G., Eddershaw, P. J., Tarbit, M. H. and
Goldfarb, P. S. (1999) Molecular modelling of the human cytochrome P450
isoform CYP2A6 and investigations of CYP2A substrate selectivity.
Toxicology 133, 1-33
2
Lewis, D. F. V. (2001) Guide to Cytochromes P450 Structure and Function.
Taylor and Francis Inc., London
3
Harrelson, J. P., Henne, K. R., Alonso, D. O. and Nelson, S. D. (2007) A
comparison of substrate dynamics in human CYP2E1 and CYP2A6. Biochem
Biophys Res Commun 352, 843-849
4
Koop, D. R., Laethem, C. L. and Tierney, D. J. (1989) The utility of pnitrophenol hydroxylation in P450IIE1 analysis. Drug Metab Rev 20, 541-551
5
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J., Gonzalez, F. J. and
Miners, J. O. (1993) Validation of 4-nitrophenol as an in vitro substrate probe
for human liver CYP2E1 using cDNA expression and microsomal kinetic
techniques. Biochem Pharmacol 46, 1975-1981
6
Peter, R., Bocker, R., Beaune, P. H., Iwasaki, M., Guengerich, F. P. and
Yang, C. S. (1990) Hydroxylation of chlorzoxazone as a specific probe for
human liver cytochrome P-450IIE1. Chem Res Toxicol 3, 566-573
7
Lucas, D., Ferrara, R., Gonzalez, E., Bodenez, P., Albores, A., Manno, M.
and Berthou, F. (1999) Chlorzoxazone, a selective probe for phenotyping
CYP2E1 in humans. Pharmacogenetics 9, 377-388
8
Kharasch, E. D., Thummel, K. E., Mhyre, J. and Lillibridge, J. H. (1993)
Single-dose disulfiram inhibition of chlorzoxazone metabolism: a clinical
probe for P450 2E1. Clin Pharmacol Ther 53, 643-650
9
Draper, A. J., Madan, A. and Parkinson, A. (1997) Inhibition of coumarin 7hydroxylase activity in human liver microsomes. Arch Biochem Biophys 341,
47-61
10
Le Gal, A., Dreano, Y., Gervasi, P. G. and Berthou, F. (2001) Human
cytochrome P450 2A6 is the major enzyme involved in the metabolism of
three alkoxyethers used as oxyfuels. Toxicol Lett 124, 47-58
97
11
Sanwald, P., David, M. and Dow, J. (1996) Characterization of the
cytochrome P450 enzymes involved in the in vitro metabolism of dolasetron.
Comparison with other indole-containing 5-HT3 antagonists. Drug Metab
Dispos 24, 602-609
12
Beulz-Riche, D., Grude, P., Puozzo, C., Sautel, F., Filaquier, C., Riche, C.
and Ratanasavanh, D. (2005) Characterization of human cytochrome P450
isoenzymes involved in the metabolism of vinorelbine. Fundam Clin
Pharmacol 19, 545-553
13
Hasegawa, A., Yoshino, M., Nakamura, H., Ishii, I., Watanabe, T., Kiuchi, M.,
Ishikawa, T., Ohmori, S. and Kitada, M. (2002) Identification of inhibitory
component in cinnamon--O-methoxycinnamaldehyde inhibits CYP1A2 and
CYP2E1. Drug Metab Pharmacokinet 17, 229-236
14
Lewis, D. F. and Hlavica, P. (2000) Interactions between redox partners in
various cytochrome P450 systems: functional and structural aspects. Biochim
Biophys Acta 1460, 353-374
15
Koop, D. R. (1986) Hydroxylation of p-nitrophenol by rabbit ethanol-inducible
cytochrome P-450 isozyme 3a. Mol Pharmacol 29, 399-404
16
Phillips, I. R. S., E.A. (2005) Cytochrome P450 Protocols: Methods in
Molecular Biology. In Spectrophotometric Analysis of Human CYP2E1Catalyzed p-Nitrophenol Hydroxylation (Chang, T. K. H. C., C.L.; Waxman,
D.J., ed.), Humana Press Inc., Totowa
17
Reinke, L. A. and Moyer, M. J. (1985) p-Nitrophenol hydroxylation. A
microsomal oxidation which is highly inducible by ethanol. Drug Metab Dispos
13, 548-552
18
Duescher, R. J. and Elfarra, A. A. (1993) Determination of p-nitrophenol
hydroxylase activity of rat liver microsomes by high-pressure liquid
chromatography. Anal Biochem 212, 311-314
19
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J. and Miners, J. O. (1993)
High-performance
liquid
chromatographic
assay
for
4-nitrophenol
hydroxylation, a putative cytochrome P-4502E1 activity, in human liver
microsomes. J Chromatogr 616, 73-78
98
20
http://www.cancer.gov/Templates/drugdictionary.aspx?CdrID=577391
(2008) Definition of Chlorzoxazone. National Cancer Institute
21
Larson, J. R., Coon, M. J. and Porter, T. D. (1991) Purification and properties
of a shortened form of cytochrome P-450 2E1: deletion of the NH2-terminal
membrane-insertion signal peptide does not alter the catalytic activities. Proc
Natl Acad Sci U S A 88, 9141-9145
22
Collom, S. L., Laddusaw, R. M., Burch, A. M., Kuzmic, P., Perry, M. D., Jr.
and Miller, G. P. (2008) CYP2E1 substrate inhibition. Mechanistic
interpretation through an effector site for monocyclic compounds. J Biol Chem
283, 3487-3496
23
Zerilli, A., Ratanasavanh, D., Lucas, D., Goasduff, T., Dreano, Y., Menard, C.,
Picart, D. and Berthou, F. (1997) Both cytochromes P450 2E1 and 3A are
involved in the O-hydroxylation of p-nitrophenol, a catalytic activity known to
be specific for P450 2E1. Chem Res Toxicol 10, 1205-1212
99
Chapter 5
Conclusions
The goal of this project was to determine the structural foundation for
the substrate selectivities of the CYP2A versus CYP2E subfamily. Because
these enzymes metabolize both common, as well as unique, small molecule
substrates, it is likely that only few key residue-substrate interactions are
responsible for those metabolic capabilities that differ between them.
Identifying these interactions could help predict the metabolism of drugs,
procarcinogens, and other xenobiotics.
A series of mutated CYP2E1 proteins were characterized for their
ability to hydroxylate the reportedly selective CYP2E1 marker substrates pnitrophenol (pNP) and chlorzoxazone (CZN) [1-5]. None of the seven
functionally active 2E1 mutants showed significant differences in activity from
the CYP2E1 wild type enzyme. However, in contrast to previous literature
reports [6-8], both CYP2A6 and CYP2A13 were observed to metabolize both
pNP and CZN with catalytic efficiencies greater than or equal to CYP2E1
(Table 5.1).
These unexpected activities of the CYP2A enzymes with
CYP2E1 substrates demonstrate that the human CYP2A and CYP2E
enzymes are more functionally similar than previously believed.
100
Table 5.1: pNP and CZN kinetic parameters for CYP2E1, CYP2A13, and CYP2A6.
CYP450
p-Nitrophenol
Km
kcat
kcat/Km
(min-1)
(µM)
(µM-1min-1)
2E1
15.9 ± 0.4
75.8 ± 4.4
2A13
30.3 ± 1.3
2A6
52.6 ± 2.4
Chlorzoxazone
Km
kcat
kcat/Km
(min-1)
(µM)
(µM-1min-1)
0.21
5.1 ± 0.4
105.5 ± 22.9
0.05
62.7 ± 7.4
0.48
22.7 ± 1.1
64.8 ± 10.4
0.23
135.8 ± 13.7
0.39
4.7 ± 0.3
100.7 ± 15.1
0.05
101
In support of our findings, Fukami et al. [9] very recently reported substantial
pNP and CZN activity by CYP2A13, however with some variations in their results
from ours. For the p-nitrophenol hydroxylation, Fukami et al. reported similar values
for CYP2E1 and CYP2A6, while CYP2A13 had a Km 6-fold lower and a kcat 2-fold
higher than the other two isozymes, thus indicating a much higher efficiency of
catalysis by CYP2A13. Although, both laboratories observed the highest efficiency
of pNP metabolism by CYP2A13, our results indicated a higher catalytic efficiency
of catalysis for CYP2A6. For the chlorzoxazone hydroxylation, Fukami reported an
8-fold higher kcat for CYP2A13 and CYP2E1 compared to CYP2A6. For the Km
values, CYP2A13 had the lowest value that was 4-fold less than that of CYP2A6,
which was 2-fold less than CYP2E1. However, the catalytic efficiency for each
enzyme matches up nicely with my finding, with similar values for both CYP2E1 and
CYP2A6, approximately 6-fold lower than that of CYP2A13. The reported variations
are likely due to several differences in the proteins and reaction conditions in these
two studies. In the Fukami report, reactions were carried out in 1.) E. coli cells, 2.)
with differing ratios of CYP450 to reductase, 3.) in the absence of cytochrome b5, 4.)
in the absence of an anti-oxidant (e.g. ascorbic acid, catalase, or SOD), and 5.) the
pNP buffer was adjusted to a pH of 7.5 versus 6.8. Their molar ratios of reductase
to CYP450 were 1.3 (CYP2E1), 2.9 (CYP2A13), and 5.8 (CYP2A6). The varying
ratios are most likely due of the relative expression of the protein in each batch of E.
coli cells containing the individual isozyme.
102
During the latter stages of the writing of this thesis, colleagues P.
Porubsky and K. Meneely determined the first crystal structures of CYP450
2E1. This recent advance has allowed us the opportunity to attempt an
explanation of the lack of effects on substrate metabolism by the CYP2E1
mutations in the substrate recognition sites (SRS) 1 and 2.
By comparing an overlay of the crystal structures of CYP2A13 and
CYP2E1, the differences in the overall structure of the two variable substrate
recognition sites 1 and 2 can be observed (Figure 5.1). The most notable
difference between these two enzymes is the longer B′ α-helix in the
CYP2A13 enzyme, which encompasses the SRS-1 region. Four amino acid
residues (107-110) extend the length of the B′ helix in CYP2A13 from that of
CYP2E1. However, there does not seem to be a significant difference in the
overall geometry and shape of the two CYP450 enzymes in this region. In the
SRS-2 region, the differences between the amino acid sequences do not
seem to have a significant impact on the overall conformation of the protein
backbone in this region.
103
Figure 5.1: An overlay of the crystal structures of CYP2E1 (tan) and CYP2A13 (light
blue). The two SRS regions 1 and 2 are highlighted in purple (CYP2A13) and orange
(CYP2E1).
I
F´
G´
B´
104
In the SRS-1 region (Figure 5.2), the altered residue L103 does not
appear to have significant structural interactions. However, it is interesting to
note that the corresponding glutamine residue in CYP2A13 is involved in
hydrogen bonding with the backbone of D107. Altering the amino acid to a
glutamine could have caused an unfavorable interaction destabilizing the
enzyme. The amino acid residue P104 was exchanged for an alanine and
appeared to have no effect on the metabolic activity of CYP2E1. This is
understandable because it appears that the proline is on the protein surface
and has very few key interactions with the rest of the protein. It points away
from the active site. The amino acid residue A105 side chain is directed into a
cleft between helices F and G. It appears there would be sufficient space in
this cleft to accommodate the additional atoms of a threonine residue, without
the disruption of the surrounding structure. The amino acid residue H107 is
located on the surface and appears to take on two conformations in the
CYP2E1 structure. The first conformation allows the formation of a salt bridge
with the side chain of the amino acid residue D102, and possible packing
interactions with P104. In the second conformation, the histidine residue
projects away from the enzyme into the surrounding solvent. The residue
H109 is involved in important hydrogen bonding with the side chain of residue
D295, effectively anchoring the B′ helix to the I helix. Mutation to a
phenylalanine would disrupt this stabilizing feature. Because the B′ helix is a
105
Figure 5.2: A view of the CYP2E1 SRS-1 looking from the heme. Large blue letters
indicate relevant helices. Residues that were exchanged are in orange sticks and the
heme is indicated in red sticks.
B´
A105
I
H109
P104
L103
H107
D295
D111
R112
106
flexible feature of the CYP450 active site, most likely moves to modulate entry
and exit from the active site and overall usually has few direct interactions
with the rest of the protein, this interaction between residue H109 and the I
helix could be critical in stabilizing the protein. The amino acid residue D111
appears to hydrogen bond with the backbone of residues G119 and N118.
However, this interaction must not be critical in the overall stabilization of the
protein since the mutation to glycine did not seem to affect the functionality of
the protein. The inability to isolate the CYP2E1 mutant enzyme R112Y in the
active form was most likely caused by the stabilizing effect of arginine’s
participation in hydrogen bonding with the amino acid residue D287. This
hydrogen bond also contributes to the anchoring of the B′ region to the I helix.
The CYP2E1 mutant protein W122A was not purified in a functional state.
This residue participates in a key interaction with the heme group. The
tryptophan is capable of hydrogen bonding to the propionic group of the heme
as well as to residue R435, which also hydrogen bonds to the propionic
group.
The first mutation made in the SRS-2 region (Figure 5.3) was at the
amino acid residue P213. It appears that this residue had little effect on the
metabolic activity of CYP2E1 because it too is positioned away from the
active site and projects into the surrounding solvent. The residue W214
participates in a hydrogen bond with amino acid residue H232. Because this
107
enzyme was purified as a functional protein, this interaction must not play a
crucial role in the overall stability of the protein. The lack of influence on the
metabolic activity was likely due to its location on the surface of the enzyme.
CYP2E1 with mutation L215G was in the inactive P420 state. This could be
due to an increase in flexibility afforded by the glycine residue, which was
detrimental to the stabilization of the enzyme. The amino acid residue P222
seems to also have little in the way of key protein interactions. Changing the
residue to a serine most likely had little effect on protein stability or substrate
metabolism because it is not oriented towards the active site, but in the
direction of the surrounding solvent.
108
Figure 5.3: CYP2E1 mutants located in the SRS-2 region. Large blue letters indicate
relevant helices. Mutated residues shown in orange with side chains as stick
representation. The heme is shown in red sticks.
G
G´
F
W214
L215
P222
F´
I
P213
109
In conclusion, the seven CYP2E1 mutants that could not be isolated as
functional protein may have been the result of detrimental effects on protein
stability caused by the exchange of those amino acid residues. Of the seven
CYP2E1 mutants purified as functional CYP450 protein no significant
changes were observed in metabolic activity. This is likely due to their
orientation away from the active site so that they do not participate in key
protein-substrate interactions.
Now that the crystal structure of CYP2E1 has been determined,
residues that modulate protein stability and substrate metabolism can be
selected rationally to determine the functional differences between these two
CYP450 subfamilies. Finally, the identification of a more selective substrate
for CYP2E1 will be needed to determine the presence and activity of CYP2E1
in microsomal preparations.
110
References
1
Tassaneeyakul, W., Veronese, M. E., Birkett, D. J., Gonzalez, F. J. and
Miners, J. O. (1993) Validation of 4-nitrophenol as an in vitro substrate probe
for human liver CYP2E1 using cDNA expression and microsomal kinetic
techniques. Biochem Pharmacol 46, 1975-1981
2
Peter, R., Bocker, R., Beaune, P. H., Iwasaki, M., Guengerich, F. P. and
Yang, C. S. (1990) Hydroxylation of chlorzoxazone as a specific probe for
human liver cytochrome P-450IIE1. Chem Res Toxicol 3, 566-573
3
Lucas, D., Ferrara, R., Gonzalez, E., Bodenez, P., Albores, A., Manno, M.
and Berthou, F. (1999) Chlorzoxazone, a selective probe for phenotyping
CYP2E1 in humans. Pharmacogenetics 9, 377-388
4
Koop, D. R., Laethem, C. L. and Tierney, D. J. (1989) The utility of pnitrophenol hydroxylation in P450IIE1 analysis. Drug Metab Rev 20, 541-551
5
Kharasch, E. D., Thummel, K. E., Mhyre, J. and Lillibridge, J. H. (1993)
Single-dose disulfiram inhibition of chlorzoxazone metabolism: a clinical
probe for P450 2E1. Clin Pharmacol Ther 53, 643-650
6
Zerilli, A., Ratanasavanh, D., Lucas, D., Goasduff, T., Dreano, Y., Menard, C.,
Picart, D. and Berthou, F. (1997) Both cytochromes P450 2E1 and 3A are
involved in the O-hydroxylation of p-nitrophenol, a catalytic activity known to
be specific for P450 2E1. Chem Res Toxicol 10, 1205-1212
7
Reinke, L. A. and Moyer, M. J. (1985) p-Nitrophenol hydroxylation. A
microsomal oxidation which is highly inducible by ethanol. Drug Metab Dispos
13, 548-552
8
Koop, D. R. (1986) Hydroxylation of p-nitrophenol by rabbit ethanol-inducible
cytochrome P-450 isozyme 3a. Mol Pharmacol 29, 399-404
9
Fukami, T., Katoh, M., Yamazaki, H., Yokoi, T. and Nakajima, M. (2008)
Human Cytochrome P450 2A13 Efficiently Metabolizes Chemicals in Air
Pollutants: Naphthalene, Styrene, and Toluene. Chem Res Toxicol 21, 720725
111