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 Km kcat/Km kcat Km kcat/Km (min-1) (µM) (µM-1min-1) (min-1) (µM) (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 O- hydroxylation 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 44 Introduction 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 87 4-5. 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 Page 1-1. 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. 69 4-1. Comparison of p-nitrophenol activity determined by visible colorimetric and HPLC UV detection methods 87 4-2. p-Nitrophenol kinetic parameters determined for CYP2E1, 2A6, and 2A13 89 4-3. Chlorzoxazone kinetic parameters determined for CYP2E1, 2A6, and 2A13 93 4-4. Comparison between CYP2E1 mutant protein activities for p-nitrophenol and chlorzoxazone 96 5-1. p-Ntirophenol and chlorzoxazone kinetic parameters for CYP2E1, 2A13, and 2A6. 101

ix

List of Schemes

Schemes Page 1.1. The catalytic cycle of cytochromes P450 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 S- oxidation, 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 11- hydroxylation 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 A B e c n a b r so b A

Figure 1.2: The heme moiety of the CYP450 enzymes with carbon monoxide complexed to the reduced iron.

O+

C-

N 2+ N! Fe O N N OH

S- OH

O cys

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 reg ions are highlighted by color: SRS-1 (yellow), SRS-2 (orange), SRS-3 (pink), SRS-4 (green), SRS-5 (blue), and SRS-6 (purple).

G´ D F´ G A´ F

B´ E I A K

J L H

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 Typical Known Localization Reaction Inducers 2A6 Liver, lung, and ER Coumarin Phenobarbital some 7-hydroxylation Barbiturates extrahepatic Dexamethasone sites Rifampin 2A7 ER 2 2 2A13 Respiratory ER Activation of NNK 2 Tract 2B6 Liver, lung ER (S)-Mephenytoin Phenobarbital N-demethylation Nicotine 2C8 Liver ER Taxol Phenobarbital 6α-hydroxylation Rifampin 2C9 Liver ER Tobutamine methyl Phenobarbital hydroxylation Rifampin Dexamethasone 2C18 Liver ER 2 Phenobarbital 2C19 Liver ER (S)-Mephenytoin Phenobarbital 4′-hydroxylation Refampin 2D6 Liver ER1 Debrisoquine Nicotine 4-hydroxylation 2E1 Liver, lung, ER Chlorzoxazone Acetone, Ethanol, other tissues 6-hydroxylation Isoniazid, Nicotine 2F1 Lung ER 3-Methylindole 2 activation 2J2 Lung ER Arachidonic 2 acid oxidations 2R1 2 2 2 2 2S1 Lung ER 2 2 2U1 2 ER 2 2 2W1 2 ER 2 2 1 Mainly ER, some detected in mitochondria. 2 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 medium- to-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

N N O N

fadrozole cotinine

O CN Cl OH

Cl F

O O Cl F O methoxyflurane 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 phenacetin styrene

N O N

naphthalene cotinine 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.

NO 2 O

HO N H

paracetamol (acetaminophen) OH

p-nitrophenol

NH2 O N N

Diethylnitrosamine aniline benzene

O

OH OH N Cl 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. In 1913, Leonor Michaelis and Maud Menten introduced the

V max[S] equation Vo = [87]. The equation describes the hyperbolic Km + [S]

!

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 Michaelis-

Menten 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.

Vmax[S] Vo = Km Vo = Vmax

) n i m /

M ! µ

. g . e ( o V 1/2 Vmax y, t

ci o l e v

l a i t

i n I

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) Heme- Containing 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, 493- 509 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 P- 450 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, 121- 131 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, 35630- 35637 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 7- hydroxycoumarin 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)-1- butanone 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 CYP450- expressing 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 tobacco- specific 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 tobacco- specific 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 substrate- enzyme 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 N- terminal 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 p- nitrophenol 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-β-

D-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 X-

Gal 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. Membrane- bound 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 (Ni-

NTA) 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 Stage 3 Stage 4 Stage 5

Equilibrate Load Wash Elution Regeneration

H H I I H I I 2+ Ni 2+ 2+ 2+ H H Ni H I Ni I 2+ Ni 2+ H Ni Ni 2+ 2+ 2+ Ni 2+ Ni 2+ Ni Ni Ni 2+ Ni Ni2+ 2+ H H Ni 2+ 2+ 2+ 2+ Ni 2+ Ni Ni I Ni Ni 2+ 2+ Ni 2+ Ni 2+ Ni Ni I 2+ 2+ H Ni Ni I H I 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 Stage 3 Stage 4 Stage 5

Equilibrate Load Wash Elution Regeneration

+ + + + + + + + + + + + +

+ + + + + + + + + + + +

+ = 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 1 minute/kb of 68 oC 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 round- bottom tubes (15 mL). The cells were then allowed to recover for 1 hour at 37 oC, 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 LB- ampicillin 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 SIZE OF DNA FRAGMENTS (bp) RESTRICTION REACTION 2E1 ENZYME CONDITIONS WILD TYPE MUTANT MUTANTS Buffer D + BSA, 6038 6038 L103Q BstE II* 1 hr at 60 oC circular linear Buffer D + BSA, 6038 6038 P104A BstE II* 1 hr at 60 oC circular linear Buffer D + BSA, 6038 6038 A105T BstE II* 1 hr at 60 oC circular linear NEBuffer 2, 1 hr 6038 H107D Bsm I** 4529, 1509 at 65 oC linear NEBuffer 2, 1 hr 6038 H109F Bsm I** 4529, 1509 at 65 oC linear Buffer C + BSA, 6038 6038 D111G Sac II* 1 hr at 37 oC circular linear NEBuffer 4, 1 hr 98, 1521, 98, 2134 R112Y Fsp I** at 37 oC 2134, 2285 3806 NEBuffer 4, 1 hr 191, 678, 191, 678, W122A Sfc I** at 37 oC 5169 2227, 2942 Buffer H + BSA, 6038 6038 P213S Pst I*** 1 hr at 37 oC circular linear Buffer H + BSA, 6038 6038 W214T Pst I*** 1 hr at 37 oC circular linear Buffer D + BSA, 6038 578, 5459 L215G Nco I* 1 hr at 37 oC linear NEBuffer 3, 1 hr 1428, 2052, 666, 1428, P222S Bpm I** at 37 oC 2558 1892, 2052 NEBuffer 4, 1 hr 19, 692, 19, 92, SRS1 rxn 1 Dra I** at 37 oC 5327 1913, 3414 NEBuffer 2, 1 hr 6038 4529, 1509 SRS1 rxn 2 Bsm I** at 65 oC linear NEBuffer 2, 1 hr 4529, 1509 6038 SRS1 rxn 3 Bsm I** at 65 oC 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 oC. 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

e d

r p n r

y e o e t o o o o o o o o o o o o o o o c

m m i e N N N N N N N N N N N N N N N d i l r s i D P

w e e h

h T

t

g g .

e e e e n n

f

r t t t t

o o g g g g g g g t o . k k r r a a a a

t t n n n n n n n r r r r s a a S s S S o o o o o o o

e e e e n e e . r r r r r r r

n d d d d t t t t t t t o c y y i o o o o o W W r r t S S S S S S S e i t e e M M M M a S t a V V

. u c i e f i t m i d

s

o

1 7 0 7 0 6 7 4 7 4 2 2 2 8 5 8

n ) 6 7 5 7 1 2 9 6 2 6 6 2 8 8 7 m E o C ......

i % t 6 9 7 9 2 2 9 6 5 7 7 2 6 1 8 s ( G c P2 6 6 6 6 6 5 4 4 4 4 4 4 4 5 4 e i t r Y t o s C n e

e r

e

d 1 6 9 6 7 6 8 0 2 8 1 4 0 3 0 a h

) ...... t m e C s 2 3 2 3 0 0 8 9 9 7 8 9 2 9 5

o T u f ( 8 8 8 8 8 8 7 7 7 7 7 7 8 7 8 e i d o f i i

s d h

e o r c )

s a m r d h i e

t e a at

r 6 3 7 3 7 4 9 5 2 2 2 5 4 4 1 g p f

h o n 3 3 3 3 3 4 4 4 4 4 4 4 6 5 4 o t l e e

s o L a n c

b o ( i h t ation c a

r

I I a

I I I I I

I I I e e I I I I I I I u o t a r c I I c

E p l t t f m r s c E E E

m m m c a d n s t t t s s s a S

s s s D N G s s s

e S

. F o P P

B

B B B i r

l

l B B B

n l l l l e n l t

l

l l l l a e a l l l a a a a a a c h a t a a a n a w

a a a n i t n n l n n n v n n n v r o n n n s o o o o o a o i

o t i o i i o o o i i

o i t o o o t i i i t t t e t i h i i i t i i i t t t s i n i i t t t t e i i i m i i i i d m d d s d t d e d d d d d o i e

d d d d e d d

d d d d d d d R s d d d n R e A A A d A R A A A A A A A A A e c e d n s

e u

T u C A

C T G A T q T C G s A C C A Red C T T G A e C e

T T C T

. T s A C C d C C

T A C T i

n A T T A t T C T T A G e A C T o T A T o A i T T T T d G t T C C C TTTTT C i e C

T C A T A l t T a A A G A C G A t A T A C G A c T o A T T C T G C A u T T G C G u A A e A T T l A T T A A A G n G G G C G m T A c T A A A A A T

C o T T A T G u C G C C A A C d A C C g A A A T G i A n T T T C G G T l C A A C C G T A re T o T A T A T T o i A C T C G T C T T C g A T T C s T i G

C T C T G C A G C T T T G e G o T C T A C T ol T T A G G C d G G A

C A G C w

A G C C A C t G G C A e G C C C e T G A A C G G

A G h C G A G C C h C C T C t A C t e G C A C T A C A C

C A G C C T T C h f C T G A C C o C C t T G G T C C G t o G T T G G G C

A G A

A T A A G T T G f C C y C n C C T G G C G G G G A r o G G G o T T T C C C A i C C G G a GC

G t A C C C t T i A T G C T T T G e A T n C C C s T G A A C A T

A AC n T A A e A C o C T A A C C A o ) C A CCTG

p ! G G C T T C G A C m C

C T A T T T 3 G A A r A C G e C

C e T T CCC C l C G G A A T o G A T G o h G T C C A f p t T T t C G G C T T T

G C GGCAGAGG T A A C

T T ! A A A CCCGC A m G s G A T A s A C C G G T T 5 G T o A G G C T A e C ( C T e T T C C

G G t

c C T A A A C C A G C c T T T G A A e G

o C C C G C C T G A A n G C C C c s A C C G n C A C T G G G T A e C C C C T T G n a C e G A A A T A G T T T C G G u e C A T C C

d G C w G T G

G A A C G G q C u A A G G C

A C T C G G e A A C C T G A A T T T G e G q e A C C A C u C C G G G C C C G G e T T C C G T T C C l T d T S i t S T C C G C G T C A G G C C C G G G G C T G

B :

o .

3 e

.

e

l

n n n n c 3 c T A F T Y A D S S G Q G x x x o n

u 4 r r r 2

9 2 5 i 4 7 3 2 1

3 5 t e 1 n 2 e

0 1 0 0 1 1 1 0 1 2 1 0 1 1 2 3 a u 2 1 1 1 1 1 1 2 2 1 S S S t 1 2 bl q u R R R A L P H P L P R H D W W a e S S S M T oligo s 65

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 SDM Dpn1 Transformation Control Mutagenic Sequencing Mutant (50ng Digestion into XL-1 Blue (Ratio of Efficiency Results and Super White:Blue (# mutant 20ng) Competent colonies) colonies: # Cells colonies (50ng/20ng) screened) 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 + W122A √ √ ***/*** 1:8 2:3 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 √ √ ****/*** 1:15 3:4 Okay rxn3

†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 Estimated Yield 2E1 Protein Absolute Purity (nmol CO Diff CYP450/ (% P450) Spectrum (A ) (nm) 420/280 1.25 L E. coli) 2E1 417 0.84 249 90%

L103Q 425 N/A 53 0% Expression #1 L103Q 425 1.09 195 10% Expression #2 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% Expression #1 SRS-1 399 N/A 93 0% Expression #2 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%

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, 15-

17], although the metabolite can also be separated from the mixture by high- pressure 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 NO2

λmax = 250 nm λmax = 250 nm

+ NaOH + NaOH

O-Na+ O-Na+

O-Na+

NO2 NO2

λmax = 405 nm 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 high- pressure liquid chromatography (HPLC).

77

Figure 4.2: The 6-hydroxylation of chlorzoxazone catalyzed by CYP450 2E1.

Chlorzoxazone 6OH-Chlorzoxazone

Cl H H N Cl N CYP450 O O - + 2e + O2 O HO O

λmax = 287 nm 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 UV-

2501PC 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 oC. 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 6- hydroxylation 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 (nmol/min/nmol) HPLC UV 10.15 ± 1.24 16.84 ± 1.55 23.28 ± 2.36 (nmol/min/nmol)

Figure 4.4: Comparison of Michaelis-Menten kinetics determined for CYP2E1 by both colorimetric and HPLC UV detection methods.

Colorimetric Assay HPLC/UV Assay

) 12 10.0 ) 10 7.5 8 5.0 6

Day 1

nmol/min/nmol nmol/min/nmol KM 44.9 ± 19.7 µM 4 KM 98.2 ± 16.2 µM 2.5 k 13.0 ± 2.6 M-1min-1 -1 -1

cat µ 2 kcat 17.3 ± 1.5 µM min Activity ( Activity 0.0 ( Activity 0 0 25 50 75 100 125 0 25 50 75 100 125 150 175 pNPpNP Concentration ((µµM)M) pNP ConcentrationConcentration ((µµM)M)

) 7.5

) 12 10 5.0 8

6 nmol/min/nmol

Day 2 nmol/min/nmol 2.5 KM 23.5 ± 8.5 µM 4 KM 146.9 ± 36.1 µM -1 -1 -1 -1

kcat 6.9 ± 0.8 µM min 2 kcat 16.8 ± 2.1 µM min Activity ( Activity

0.0 ( Activity 0 0 25 50 75 100 125 0 50 100 150 200 250 300 pNP Concentration (µM) pNP Concentration (µM) 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 CYP450 -1 -1 -1 (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

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 3- fold 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 CYP450 -1 -1 -1 (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

Figure 4.7: An overlay of the enzyme kinetic parameters of CZN metabolism by CYPCYP4502E1, CYP2A 6CZN, and C YMetabolismP2A13.

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 p- nitrophenol 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 7- hydroxylase 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 CYP2E1- Catalyzed 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 p- nitrophenol (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.

p-Nitrophenol Chlorzoxazone

CYP450 kcat Km kcat/Km kcat Km kcat/Km (min-1) (µM) (µM-1min-1) (min-1) (µM) (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 0.05

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.

A105 B´ I

P104 H109

L103 H107

D295

R112 D111

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 F

G´ W214

L215 I P222

F´ 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 p- nitrophenol 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, 720- 725

111