Department of Drug Metabolism and Pharmacokinetics,
Rhône-Poulenc Rorer, Collegeville, Pennsylvania (J.C.S., R.B.W.);
Department of Pharmacology and Toxicology, University of Texas Medical
Branch, Galveston, Texas (T.L.D., J.R.H.); Department of Structural
Analysis, Rhône-Poulenc Rorer, Collegeville, Pennsylvania (E.O.);
Selectide Corporation, Tucson, Arizona (G.R.H.)
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Introduction |
The
cytochrome P-450 (CYP) superfamily is the primary catalyst of the
oxidative biotransformation of therapeutic agents. The role of the
CYP3A subfamily in drug metabolism has been extensively studied,
primarily due to the ability of CYP3A forms to metabolize numerous
drugs across several therapeutic classes. CYP3A4 is characterized by
high levels in human liver (an average of 28% of total hepatic CYP
levels). Although CYP3A4 is not polymorphically expressed (Guengerich,
1995
), the wide range of enzyme levels in patients can underlie inter-
and intra-individual variability in clinical pharmacokinetics and
efficacy. In contrast, CYP3A5 clearly displays a genetic polymorphism
(Wrighton et al., 1989) that has been linked to a point mutation at
exon 11 (Jounaidi et al., 1996).
CYP forms have been implicated in the sequential oxidation of sulfur
containing compounds to sulfoxide and sulfone metabolites. RPR 106541 {20R-16
,17-[butylidenebis(oxy)]-6,9-difluoro-11
-hydroxy-17-(methylthio)androsta-4-en-3-one, Fig. 1} is an airway-selective
17-thiosteroid currently in preclinical development for the treatment
of asthma. Similar steroid derivatives have been proven to be effective
against the allergic inflammatory responses of the lung (Check and
Kaliner, 1990). These compounds undergo extensive first-pass
biotransformation, thus limiting systemic exposure and the potential
for suppression of the hypothalamo-pituitary-adrenal axis (Jonsson et
al., 1995).
The initial focus of this work was to identify the primary route(s) of
oxidation of RPR 106541 in human liver microsomes. The involvement of
individual CYP forms in the production of two sulfoxide enantiomers was
then assessed via correlations with marker CYP activities, selective
chemical and biological inhibitors, and expressed CYP forms. The second
objective was to use apparent advantages of the RPR 106541 structure in
concert with site-directed mutagenesis of SRS residues to study the
active site of CYP3A4. The use of site-directed mutagenesis to analyze
putative CYP substrate recognition site (SRS) residues has been
successful in identifying key active site residues for CYP2D6 (Ellis et
al., 1995), CYP2C forms (von Wachenfeldt and Johnson, 1995), and
several CYP2B forms (Halpert and He, 1993; Hasler et al. 1994).
Hydroxylation profiles of endogenous steroids such as progesterone,
androstenedione, and testosterone have been used for structure-function
studies with CYP3A4 (Harlow and Halpert, 1997; Szklarz and Halpert,
1997; Domanski et al., 1998). However, these compounds are less than optimal due to properties of the steroid and the large, accommodating active site of CYP3A4. Specifically, the strong electronic effects of
the 6-carbon are presumed to favor hydroxylation at this site by CYP3A
forms, thus limiting the role of enzymatic constraints in determining
product profiles (Harlow and Halpert, 1997). With RPR 106541, a
fluorine atom blocks oxidation at the 6- site. Other sites of
endogenous steroid oxidation by CYP3A enzymes include the 2-, 15-, and
16-carbon positions; however, with RPR 106541, the
16,17-butylidenedioxy substituent prevents metabolism at the 16-position and is likely to sterically hinder attack at the 15-carbon position. Therefore, RPR 106541 offers the advantage of having enzyme
constraints dictate metabolism by directing oxidation to a steroid side
chain rather than the steroid nucleus. Consequently, we have studied
the effects of several CYP3A4 SRS mutants on RPR 106541 sulfoxidation
and have shown that the ratio of formation of the sulfoxide enantiomers
is sensitive to mutations at residues 210, 304-305, and 370.
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Materials and Methods |
Chemicals.
RPR 106541 (Batch TP2969 and Lot JYW
417/3), RPR 104065 (methyl 16,17-butylidene
bis(oxy)
6,9-difluoro-11,21-dihydroxy-3-oxoandrosta-1,4-diene-17-thiocarboxylate; Batch NCP80), RPR 112020 (S-diastereomer sulfoxide of RPR
106541; Batch JYW 473P), RPR 112903 (R-diastereomer
sulfoxide of RPR 106541; Batch JYW 490/2), RPR 112022 (sulfone
metabolite; Batch JYW 47P), and RPR 112023 (3-hydroxy derivative of RPR
106541; Batch JYW 47P) were all obtained from RPR, Dagenham Research
Center (Dagenham, England). CHAPS
{3-[(3-cholamidopropyl)-dimethylammonio]-1-propanesulfonic acid},
glucose 6-phosphate, NADP+, -NADPH, glucose
6-phosphate dehydrogenase, and troleandomycin (TAO) were purchased from
Sigma Chemical Co. (St. Louis, MO). Methyl-t-butyl ether and
isopropyl alcohol were purchased from EM Science (Gibbstown,
NJ). Acetonitrile was purchased from J.T. Baker (Philipsburg,
NJ). All other chemicals were obtained from standard vendors.
Biological Reagents.
Human liver samples were obtained
through organ procurement agencies (Anatomic Gift Foundation, Woodbine,
GA; International Institute for the Advancement of Medicine/In Vitro
Technologies, Exton, PA; National Disease Research Interchange,
Philadelphia, PA; and the Association for Human Tissue Users, Tucson,
AZ) in accordance with proper ethical procedures for consent. Rat
(Sprague-Dawley), dog (beagle), and human liver microsomes were
prepared according to the procedure of Wang et al. (1983). Anti-CYP3A1
IgG was purchased from Human Biologics (Phoenix, AZ), and IgG stock
solutions were diluted with PBS. Microsomes prepared from human
lymphoblastoid cells or baculovirus-infected insect cells transfected
with individual CYP forms were purchased from Gentest Corp. (Woburn,
MA). Mutagenesis of CYP3A4 residues and enzyme expression were
performed as described previously (Domanski et al., 1998).
Enzyme Assays.
The following enzyme assays were used to
monitor specific CYP forms as described previously (Heyn et al., 1996):
100 µM phenacetin O-deethylation for CYP1A2, 50 µM
coumarin 7-hydroxylation for CYP2A6, 200 µM S-mephenytoin
N-demethylation for CYP2B6, 200 µM S-mephenytoin 4'-hydroxylation for CYP2C19, 0.5 mM
chlorzoxazone 6-hydroxylation for CYP2E1, and 50 µM nifedipine
oxidation for CYP3A4. CYP2D6 activity was measured by bufuralol
1'-hydroxylation (Kronbach et al., 1987) at a substrate concentration
of 100 µM, and diclofenac 4'-hydroxylation (20 µM diclofenac) was
used as a CYP2C9 marker activity (Leemann et al., 1993).
Incubations of RPR 106541 with liver microsomes were conducted under
the following conditions: 0.65 mg/ml microsomal protein, 50 mM Tris/HCl
(pH 7.4), 1 mM NADP+, 1 U/ml glucose 6-phosphate
dehydrogenase, and 100 µM RPR 106541 in a total volume of 1 ml. For
the antibody inhibition experiments with RPR 106541, the microsomal
protein concentration was reduced to 0.12 mg/ml and the incubation time
was increased to 20 min to conserve antibody. Antibody, microsomal
protein, and all other incubation components except substrate and
glucose 6-phosphate were incubated with gentle shaking at room
temperature for 30 min before the addition of the substrate.
Incubations using CYP forms expressed in baculovirus-infected insect
cells (Gentest Corp.) used 50 pmol CYP in a total volume of 0.2 ml, 60 min incubation time, and direct precipitation of protein followed by
HPLC analysis as described below. Samples were preincubated for 3 min
at 37°C in a shaking water bath, and the reaction was started by the
addition of glucose 6-phosphate (10 mM final concentration). Reactions were terminated by the addition of 4 ml of methyl-t-butyl
ether and the samples were extracted by mechanical shaking for 15 min followed by centrifugation for 10 min at 4000 rpm. For quantitative studies, 3 ml of the organic phase were removed and transferred to a
clean tube. Samples were then evaporated with nitrogen and reconstituted in mobile phase (50% A/50% B, see below) before injection.
Samples were analyzed by HPLC using a Spherisorb ODS-1, 250 × 4.6 mm, 5-µm analytical column (Phase Separations, Franklin, MA)
with a Zorbax ODS 4 mm × 1.25 cm guard column (MacMod Analytical, Chadds Ford, PA). A gradient elution was used at 1 ml/min with mobile
phase A (30% acetonitrile/65% water/5% isopropyl alcohol) and B
(55% acetonitrile/40% water/5% isopropyl alcohol): initial conditions were 50% solvent B for the first 5 min increasing to 70% B
from 5 to 19 min, then increasing to 100% B from 22 to 26 min, and
holding at 100% B until the end of the run (34 min). A Spherisorb
ODS-1, 250 × 10 mm, 5-µm semipreparative column (Phase Separations) in combination with a 60 × 10 mm guard column packed with the identical material was used for larger scale separations. The
flow rate of 4.73 ml/min represents a linear velocity equivalent to
that of the analytical column. Kinetic analysis of the human liver
microsome experiments was performed using a one-site model (GraphPad
Software, Inc., San Diego, CA).
Mutagenesis and Initial Characterization of CYP 3A4 Residues 373, 479, and 480.
Plasmid pS3A4His (Domanski et al., 1998) was used as
the template for construction of L373A, L479F, and G480Q. The Expand PCR kit (Boehringer Mannheim, Indianapolis, IN) was used with a common
reverse primer that overlapped 21 bases with 479F and 480Q primers,
which contained the desired mutations plus a silent mutation that
removed a HindIII site (Fig.
2). After amplification, the reactions
were incubated with DpnI, an enzyme that will only digest
methylated DNA. This process removed any circular, unmutated template
plasmid. The DNA was transformed into DH5 cells, and plasmid DNA
from the resulting colonies was isolated and digested with
HindIII to screen for the desired mutations, because these constructs lacked one HindIII site. The entire 3A4 coding
region was confirmed by sequencing (University of Arizona Sequencing Facility, Tucson, AZ). Primers L373A and L373 reverse were used to
introduce the L373A mutation into the 3A4 cDNA (Fig. 2). After amplification of the desired region, the MunI to
KpnI fragment was subcloned into the pS3A4His plasmid,
replacing the wild-type fragment. The mutated region was sequenced to
verify the presence of the desired mutation. The construction of the
remaining mutants presented in Table 3 has been described previously
(Harlow and Halpert, 1997, 1998; He et al., 1997; Domanski et al.,
1998). The Expand PCR kit and DpnI were purchased from
Boehringer Mannheim. All other restriction enzymes and Taq
polymerase were purchased from Life Technologies (Grand Island, NY).
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Fig. 2.
Primers used to construct mutations as CYP 3A4
residues 373, 479, and 480. The underlined residue denotes the silent
mutation removing a HindIII restriction site. The
boldface bases denote mutations to make residue conversions.
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Initial characterization of L373A, L479F, and G480Q was performed using
protein preparations that were expressed in Escherichia coli
and purified from solubilized membrane preparations using the Talon
Affinity system (Clontech, Palo alto, CA) as described previously (Domanski et al., 1998). Testosterone and progesterone hydroxylation assays were also performed as described previously (Domanski et al., 1998).
Mutant Enzyme Assays.
CYP3A4 mutants were provided as
purified preparations (Harlow and Halpert, 1997; He et al., 1997;
Domanski et al., 1998). Preparations were reconstituted before
incubation with RPR 106541 by the addition of the following reagents
(in order of addition): CYP, 4-morpholinepropanesulfonic acid (MOPS)
buffer [100 mM MOPS (pH 7.3), 10% glycerol, 0.2 mM dithiothreitol, 1 mM EDTA], CHAPS (10% stock), dioleoylphosphatidylcholine (DOPC; 1 mg/ml stock), reductase, and cytochrome
b5. The
CYP/reductase/b5 ratio was chosen as
1:4:2 based on previous data showing this ratio was optimal for
testosterone and progesterone hydroxylase activities (Domanski et al.,
1998). The CHAPS concentration was 0.25-0.4%, and the DOPC
concentration was 0.1 mg/ml. The MOPS buffer was added only to dilute
all the CYP samples to an equal concentration before adding the
remaining reagents. The reconstitution was carried out at room
temperature for 10-15 min. The reaction volume ranged from 666 µl to
1.040 ml with the following conditions: 50 pmol CYP, 100 µM RPR
106541, 50 mM HEPES buffer (pH 7.6), 15 mM MgCl2, 0.1 mM EDTA, 0.04% CHAPS, and 0.1 mg/ml DOPC, and the reactions were
started by the addition of 1 mM NADPH (final concentration). The
incubation time was 60 min, and the reactions were terminated and
analyzed as described above.
Computer Modeling.
The model of CYP 3A4 developed previously
(Szklarz and Halpert, 1997) was used to dock substrate RPR 106541 into
the active site in the proper orientation for the formation of the
R-sulfoxide metabolite to understand the changes in
specificity resulting from certain amino acid substitutions. The
substrate was placed into a reactive binding orientation, with the
oxidation site fixed at 3.24-3.5 from the heme oxygen as described
previously (Szklarz and Halpert, 1997). This leads to sulfoxidation and
formation of the R-sulfoxide metabolite. Conformational
analysis of RPR 106541 was performed with the SEARCH COMPARE module of
INSIGHT II (Molecular Simulations, Inc, San Diego, CA), and the
total energy of the docked substrate and protein were analyzed using the DOCKING module (Szklarz et al., 1995). The docking interaction between the enzyme and the substrate was optimized using energy minimizations within INSIGHT II (Szklarz et al., 1994).
Structural Analysis.
Liquid chromatography/mass spectrometry
(LC/MS) analyses were performed on a Sciex API III spectrometer
(Sciex, Foster City, CA) interfaced to a Hewlett-Packard model 1050 chromatography system (Palo Alto, CA). The HPLC separations used
a combined gradient on a microbore column (Spherisorb ODS-1, 1 × 150 mm, 5 µm, Phase Separations). The mobile phase comprised A = 20 mM ammonium acetate/isopropyl alcohol, 90:10, and B = 20 mM
ammonium acetate/acetonitrile/isopropyl alcohol, 45:45:10. The gradient
used was 50% B for the first 5 min, then increased to 70% B over 20 min and held for 5 min, and finally increased to 100% B over 5 min.
Using MS/MS experiments, structural information was derived from
molecular fragment ions produced either by collision-induced
dissociation using IonSpray, or at higher energies with heated
nebulizer and corona discharge. Electron impact mass spectra
were acquired at 70 eV using a Finnegan (San Jose, CA) model 4500 spectrometer with a direct insertion probe. Proton NMR spectra were
obtained at 500 MHz with a Varian UnityPlus (Varian, Palo Alto, CA)
spectrometer atmospheric pressure chemical ionization with the analyte
in CDCl3 and tetramethylsilane as internal
reference. Infrared spectra were recorded at 4 cm
1 resolution on a Nicolet model 740 FTIR with
an IRPLAN microscope (Nicolet, Madison, WI).
X-Ray Crystallographic Structure Determination of RPR
112020.
A single crystal of RPR 112020 at a temperature of 153 K
was used for the x-ray crystallographic measurements on an
Enraf-Nonius CAD-4 diffractometer (Enraf-Nonius, the
Netherlands), with graphite crystal monochromatized Mo radiation.
An empirical 
-scan absorption correction was applied with the data
from nine reflections. The intensities of six standard reflections,
monitored every hour of crystal X-ray exposure time, showed no change
in the average intensity over the measurement period. Calculations were
performed using TEXSAN (Molecular Structure Corp., Molecular Structure, The Woodlands, TX) and SHELXTL (Siemens Industrial Automation, Inc.) software programs (Siemens, Munich, Germany). The
structure was solved with direct methods and refinement was by
full-matrix least-squares with anisotropic temperature factors for the
C, F, O, and S atoms and isotropic terms for the H atoms. The C-linked H atoms were positioned from the heavy atom framework; the hydroxyl H
atoms were located in a difference electron density map. There were two
crystallographically independent molecules per unit cell. The absolute
configuration was established by refinement of the Flack x
parameter (0.038 for this determination; Flack, 1989).
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Results |
Identification of RPR 106541 Sulfoxide Metabolites.
Preliminary metabolite profiling studies consisted of the incubation of
rat, dog, and human liver microsomes with RPR 106541 followed by HPLC
analysis. These incubations produced two major metabolites with
retention times of 13.1 (M1) and 13.9 min (M2;
data not shown). Preliminary LC/MS analysis of the organic extract of
incubations of RPR 106541 with dog liver microsomes showed two closely
eluting peaks that each produced a molecular ion of 473 (M + H+). Because of the highly efficient conversion
of RPR 106541 to M2 by female rat liver microsomes, larger
scale metabolite generation was undertaken using this enzyme source.
Specifically, 30 identical 60-min incubations of 3 mg/ml female rat
liver microsomal protein and 300 µM RPR 106541 were conducted.
Following extraction, the samples were pooled and injected onto a
semipreparative (250 × 10 mm) Spherisorb ODS-1 column.
M2 was collected from multiple injections, evaporated under
nitrogen, and dried in vacuo. The isolated sample was then analyzed by
infrared, NMR (proton), and mass (electron impact, LC MS/MS) spectrometry.
Figure 3 shows the electron impact
spectra of M2 from dog liver microsome incubations. The
fragment ions revealed the loss of CH3SOH
(product ion m/z 409) and
C4H7OH (product ion
m/z 337), consistent with the structure of a sulfoxide of
RPR 106541. The fragmentation patterns for M2 isolated from
rat liver microsome incubations and from the dog liver microsome
incubations were identical with each other and to the fragmentation
pattern for M1. The proton NMR spectra for M2
produced characteristic shifts in the positions of the hydrogen atoms
adjacent to the sulfur atom. Specifically, the hydrogens of the
thiomethyl group (H13) of RPR 106541 had a chemical shift of 2.134 ppm;
in contrast M2 exhibited a H13 chemical shift of 2.694 ppm.
The deshielded position (relative to RPR 106541) of the
S-methyl resonance (hydrogen position H13) for M2
is indicative of a sulfoxide. In addition, the appearance of this
resonance as a singlet at high resolution indicates that M2
consists of a single sulfoxide enantiomer. Shifts were also observed
for hydrogens at positions 3 (t, 5.616 ppm), 5 (d, 5.273, 5.262), and 23 (s, 1.106).
M2 was later shown to cochromatograph with a synthetic
standard of the S diastereomer RPR 112020 (data not shown).
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Fig. 3.
Electron impact spectra of M2 isolated
from incubations of RPR 106541 with rat liver microsomes. The
fragmentation ions show the consecutive losses of the
CH3SOH [m/z 409, (M + H)+ ] and C4H7OH [m/z 337, (M + H)+] groups, consistent with sulfoxide formation. In
combination with NMR and X-ray crystallography data, this compound was
identified as the S-sulfoxide metabolite (RPR 112020).
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Finally, a single crystal X-ray structure determination of RPR 112020 was performed to unambiguously determine the stereochemistry of the
sulfoxide position. Based on the refined Flack x parameter of 0.038, the absolute configuration of the sulfoxide was assigned as
S. By default, RPR 112903 (M1) was assigned the
R configuration.
Identification of the Human CYP Forms Responsible for RPR 106541 Sulfoxidation.
Based on preliminary kinetic experiments examining
the effect of protein concentration and incubation time on the
linearity of R- and S-sulfoxide formation, a
protein concentration of 0.65 mg/ml and a 10-min incubation time were
used. The kinetics of RPR 106541 sulfoxidation were then determined for
three human liver microsome samples (HL-04, HL-05, and HL-06). The
respective Km and
Vmax values for all three samples are
given in Table 1. For
R-sulfoxide formation, the
Km values ranged from 55 to 73 µM
and the Vmax values from 0.70 to 2.01 nmol/min/mg protein. For S-sulfoxide formation, the
Km values ranged from 21 to 59 µM
and the Vmax values from 0.84 to 2.46 nmol/min/mg protein. The three Km
values for each metabolite were compared by a two-sample t
test and determined not to be statistically different
(p 
.05). A concentration of 100 µM RPR 106541 was
thus chosen as saturating for the determination of the rates of
sulfoxidation for a panel of human liver microsome samples. The results
for 15 human liver microsome samples and the correlation with
CYP3A4-marker nifedipine oxidase activity are shown in Fig.
4A and B. For R-sulfoxide
formation, the enzyme activity ranged 5.8-fold from 0.31 to 1.81 nmol/min/mg protein, and for S-sulfoxide formation, enzyme
activity ranged 4.7-fold from 0.44 to 2.06 nmol/min/mg protein. These
activities were then correlated with CYP marker activities previously
determined for the same panel of human liver microsome samples. As
shown in Table 2, of the eight marker
activities included in the analysis, CYP3A4-catalyzed nifedipine
oxidase activity produced the best correlation with the formation of
both sulfoxides. Specifically, the correlation coefficients
(r2) were 0.84 and 0.91 for
S- and R-sulfoxide formation, respectively. The
relatively strong correlation of sulfoxide formation with CYP2D6-marker
bufuralol 1-hydroxylase activity was not supported by subsequent
studies examining the effect of the CYP2D6 inhibitor quinidine or of
expressed CYP2D6 on RPR 106541 sulfoxidation (see below).
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TABLE 1
Apparent Km and Vmax values of
RPR 106541 sulfoxidation by human liver microsomes
Incubations were conducted as described in Materials and
Methods with RPR 106541 concentrations ranging from 10 to 400 µM. Kinetic parameters were determined using a one-site model.
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Fig. 4.
Rates of formation of the R- (A) and
S-sulfoxides (B) for 15 human liver microsomes samples
correlated with the corresponding nifedipine oxidase activity. The
r2 values were calculated from linear
regression analysis.
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TABLE 2
Correlation of various CYP form-selective activities in a panel of
human liver microsomes with conversion of RPR 106541 to the
R- and S-sulfoxide metabolites
Substrate concentrations and assay conditions are given in
Materials and Methods. n = 15 for all
analyses except CYP2E1 (n = 14).
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The ability of CYP form-selective inhibitors to alter the sulfoxidation
of RPR 106541 was also examined. As shown in Fig. 5, RPR 106541 sulfoxidation was not
affected by inhibitors of CYP 1A2 (furafylline), 2C9 (sulfaphenazole),
2D6 (quinidine), or 2E1 (diethyldithiocarbamate). Coumarin, a
competitive inhibitor of CYP2A6, produced minimal inhibition of
sulfoxide formation (~15% for each diastereomer). The CYP3A4
substrate cyclosporin A produced 44% inhibition of
R-sulfoxide formation and 38% inhibition of
S-sulfoxide formation. Troleandomycin produced ~50%
inhibition of each reaction at a concentration of 50 µM, thus
implicating CYP3A forms in RPR 106541 sulfoxidation. Antibody
inhibition results for RPR 106541 sulfoxidation paralleled the chemical
inhibition data. As shown in Fig. 6, 5 mg
anti-CYP3A1 IgG/mg microsomal protein produced 64% and 70% inhibition
of S- and R-sulfoxide formation, respectively.
The same ratio of antibody inhibited 82% of CYP3A4-marker nifedipine
oxidase activity.
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Fig. 5.
Inhibition of human liver microsomal RPR 106541 sulfoxidation by CYP-form selective inhibitors. The open bars represent
R-sulfoxide formation, and the striped bars represent
S-sulfoxide formation. Abbreviations used are Sulf
(sulfaphenazole), CsA (cyclosporin A), TAO, Quin (quinidine), and DDC
(diethyldithiocarbamate). Furafylline, sulfaphenazole, TAO, and DDC
were preincubated with human liver microsomes in the presence of NADPH
before the addition of substrate. The percentage of activity remaining
is calculated based on the respective control values.
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Fig. 6.
Inhibition of RPR 106541 sulfoxidation by anti-CYP3A1
antibody. The percentage of control represents enzyme activity in the
presence of anti-CYP3A1 antibody relative to enzyme activity in the
presence of the same ratio of preimmune (rabbit) sera to microsomal
protein. The rate of R-sulfoxide ( ) and
S-sulfoxide ( ) formation is shown in comparison to
inhibition of CYP3A4 marker nifedipine oxidase activity ( ). Details
are provided in Materials and Methods.
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Finally, the sulfoxidation of RPR 106541 by a panel of microsome
samples of expressed CYP forms was measured. CYP1A2, -2B6, -2D6, -2E1,
and -2A6 did not metabolize RPR 106541. CYP2C19 and -3A4 catalyzed the
formation of the R-sulfoxide at rates of 0.91 and 1.23 nmol/min/nmol CYP, respectively. For S-sulfoxide formation, the respective rates in nmol/min/nmol were as follows: CYP3A4, 1.33;
CYP3A5, 1.21; CYP2C9, 2.27; and CYP2C19, 3.71. Because other investigators have shown a requirement for cytochrome
b5 with CYP3A-dependent reactions
(Gillam et al., 1993; Yamazaki et al., 1996), a separate experiment
examined the effect of b5 addition on
RPR 106541 sulfoxidation by CYP3A and -2C forms. The addition of 1 or 2 nmol b5/nmol CYP did not affect the
conversion of RPR 106541 to the R-sulfoxide, except that
CYP3A5 now displayed a rate of R-sulfoxide formation of 0.5 nmol/min/nmol CYP. Addition of b5 did
not affect the rate of S-sulfoxide formation by CYP3A4, but
did produce low levels (<50%) of stimulation of
S-sulfoxide formation by CYP3A5, -2C9, and -2C19 (data not shown).
Although CYP2C forms displayed the highest rates of
S-diastereomer formation, the involvement of CYP2C forms in
the metabolism of RPR 106541 was not supported by other phenotyping
approaches. Specifically, RPR 106541 sulfoxidation did not correlate
with human liver microsomal CYP2C9-catalyzed tolbutamide 4-hydroxylase activity (r2 = 0.01 and 0.00 for
R- and S-sulfoxide formation, respectively). Also, 100 µM sulfaphenazole produced only 4% inhibition of
R-sulfoxide formation and 13% of S-sulfoxide
formation. This finding is in contrast to the >80% inhibition of
tolbutamide hydroxylation reported by Newton et al. (1995) at the same
concentration of sulfaphenazole.
RPR 106541 Sulfoxidation by CYP3A4 Mutants.
The effect of
site-directed mutagenesis of various CYP3A4 SRS residues on RPR 106541 sulfoxidase activity was studied using several purified preparations.
These mutants were engineered based on a proposed CYP3A4
three-dimensional model and the localization of putative 3A4 SRSs based
on alignment of 3A4 with nonmammalian CYP sequences (Szklarz and
Halpert, 1997) as originally introduced by Gotoh (1992) for the CYP2
family. Table 3 shows the rates of
formation of each enantiomer and the ratio of R:S-sulfoxide formation for each mutant and wild-type 3A4. The wild type produced an
enantiomer ratio of 0.89, similar to that of commercially available CYP3A4 expressed in insect cells (0.92). Within SRS-2, the Leu210 
Ala mutation resulted in a sharp decrease in the sulfoxide ratio (0.24). The effect was decreased (0.55) for L211A. Substitution of Phe
for Leu at 211, however, did not alter the ratio of sulfoxide formation
compared with the wild type.
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TABLE 3
Metabolism of RPR 106541 by CYP3A4 mutants expressed in E. coli
Assays were conducted as described in Materials and Methods
section for Mutant Enzyme Assays. Values are average of
duplicate samples.
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SRS-4 is located within the highly conserved I helix and has been shown
to influence both the rate and stereoselectivity of endogenous steroid
hydroxylation (Domanski et al., 1998). With RPR 106541, SRS-4
substitutions were clearly more detrimental to formation of the
R-diastereomer in comparison to the
S-diastereomer. For I301A, the decrease in the ratio to 0.42 was entirely dependent on the drop in R-sulfoxide formation.
With F304A the ratio decreased to 0.25 due to a 3-fold decrease in
R-sulfoxide formation and a slight increase in the
S formation. S-sulfoxide formation was reduced by
the change from the -methyl at 305 to an -hydrogen, resulting in
a lower overall ratio. The substitution of valine at 305 produced both
the lowest enantiomer ratio and the lowest rate of
R-sulfoxide formation of any mutant tested (0.20).
Residues 369, 370, and 373 have been shown to dictate testosterone and
progesterone metabolite profiles (He et al., 1997). Although the
corresponding 6 and 16 sites of metabolism are not available for
hydroxylation with RPR 106541, altered sulfoxide production could
provide evidence that these residues are involved in substrate binding
within the enzyme active site. RPR 106541 sulfoxidation was not altered
by I369V relative to the wild type, and L373A showed a diastereomer
ratio close to the wild type, although with uniformly decreased rates
of R- and S-sulfoxide formation. However, A370V
showed a 50% decrease in R-sulfoxide formation whereas
S-sulfoxide formation was increased ~2-fold. This mutation
has been postulated to stabilize the 16 binding orientation of
progesterone (He et al., 1997) and may have a similar effect on the
positioning of RPR 106541 for sulfoxidation in the S configuration.
The effect of amino acid substitutions in the proposed SRS-6 of CYP3A4
on enzyme activity has not been reported. R-sulfoxide formation was selectively affected by a L479 Phe alteration as
evidenced by the 0.42 ratio. This change in activity for L479F is in
contrast to the lack of effect of this mutation on the
6-hydroxy to 16-hydroxy ratio for the metabolism of
progesterone or testosterone. (All of these mutants maintained
6-OH to 16-OH ratios similar to CYP 3A4 wild type, although G480Q
had an overall decrease in activity of ~8-fold and L373A displayed a
2-fold decrease.)
Interpretation of Mutagenesis Data Using Molecular Modeling.
Previously, the CYP-3A4 three-dimensional model has been used
successfully to interpret mutagenesis data (Harlow and Halpert, 1997,
Domanski et al., 1998). Therefore, this methodology was employed to
interpret the changes in R-sulfoxide formation displayed by
several mutants studied including L210A, I301A, F304A, A305V, A370V,
and L479F. Figure 7 clearly illustrates
and supports the results obtained in several of these instances. For
example, the increased size of the valine side chain in A305V and A370V
causes increased van der Waals overlaps, inhibiting formation of the R-sulfoxide. The same decrease in R-sulfoxide
formation was observed for mutant L479F, where a leucine is replaced
with a very bulky phenylalanine. I301A and F304A, however, have large
side chains replaced by the much smaller alanine. The positioning of
these residues suggests that the lack of the bulkier side chains may allow for excess movement of the substrate in the binding pocket, limiting residence time in a productive orientation. The side chain of
mutant L210A appears to be too far from the docked substrate to play a
significant role in substrate specificity; in fact, it is located more
than 5 Å from the docked substrate. However, this residue may be
located in the channel of entry, a possibility that cannot be addressed
with current modeling methodology.
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Fig. 7.
Substrate RPR 106541 docked into a CYP3A4 model in
the proper orientation for the formation of the
R-sulfoxide metabolite. The substrate is shown in gray,
as are the side chains of the mutated residues A210, A213, A301, A304,
V305, V370, and F479.
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Discussion |
The importance of identifying the major routes of metabolism for a
new chemical entity and the enzymes involved is well recognized. The
primary strategy for generating this information first typically involves coupling in vitro metabolism systems with structural analysis
(LC/MS, NMR, etc.) to identify the major metabolites. If the reaction
is then determined to be CYP-catalyzed, a variety of approaches
(correlation with marker activities in a bank of human liver
microsomes, chemical/antibody inhibition, expressed enzymes) are then
used to determine the responsible human CYP form(s) (Wrighton et al.,
1993; Guengerich, 1995). The experimental resources required for this
work are significant, and thus limits the number of new chemical
entities that can be evaluated as human pharmaceuticals. The
development of more empirical and predictive methods could expedite
this process and allow a description of the enzymology of a reaction to
be incorporated in the new chemical entity selection process at an
earlier stage. Specifically, methods using photoaffinity probes,
molecular modeling, chimeric enzymes, and site-directed mutagenesis
have been used to identify critical binding residues of mammalian CYP
forms or to predict the site(s) of CYP metabolism (von Wachenfeldt and
Johnson, 1995). In addition, homology models based on the crystal
structure of bacterial CYP101 that could be applied to computer aided
drug design have been generated for several human CYP forms, including
CYP2C9 (Korzekwa and Jones, 1993), CYP2D6 (Koymans et al., 1993), and
CYP3A4 (Ferenczy and Morris, 1989; Szklarz and Halpert, 1997).
In our present study, the sulfoxidation of RPR 106541 by human liver
microsomes has been characterized by using a combination of reaction
phenotyping techniques followed by an analysis of changes in metabolism
by CYP3A4 SRS mutations. The R- and S-sulfoxide diastereomers are the major oxidative metabolites formed by human liver
microsomes. The correlation of RPR 106541 sulfoxidase activities with
human liver microsomal CYP3A4 marker nifedipine oxidase activity and
the metabolism of RPR 106541 by expressed CYP3A4 and CYP3A5 strongly
suggest that CYP3A forms are the primary catalysts of RPR 106541 sulfoxidation in human liver microsomes. The involvement of expressed
CYP2C9 and CYP2C19 in RPR 106541 sulfoxidation was not supported by
correlation analysis or chemical inhibition data. Yamazaki and Shimada
(1997) have shown that CYP3A4, CYP2C9, and CYP2C19 are all involved in
the 21-hydroxylation of progesterone, a site similar in spatial
configuration to the S-methyl substituent of the D ring of
RPR 106541. The contribution of CYP2C forms to RPR 106541 metabolism
could explain why chemical inhibition studies showed that the
CYP3A-selective inhibitor TAO produced only moderate inhibition of RPR
106541 sulfoxidation (~50%). However, there is precedence for the
finding of incomplete inhibition by TAO of reactions shown to be
CYP3A-dependent by other reaction phenotyping approaches (Guengerich,
1990; Fleming et al., 1992; Kumar et al., 1994). Finally, antibody
inhibition studies show that anti-CYP3A1 antibody inhibits the
conversion of RPR 106541 to both the R- (70% of control)
and S-sulfoxides (64% of control).
The evaluation of 14 CYP3A4 mutants for RPR 106541 sulfoxidase activity
represents a comprehensive evaluation of CYP3A4 SRS sites 2, 4, 5, and
6 and a logical extension of the phenotyping studies to CYP3A4
structure-function analysis. Within SRS-2, the Leu210 Ala mutation
decreased the enantiomer ratio from 0.89 (wild type) to 0.25, predominantly due to a decrease in R-sulfoxide formation.
This substitution has been shown previously to decrease the rates of
testosterone 2- and 15-hydroxylation and to alter the response to
stimulation by -naphthoflavone, suggesting that effector and binding
sites of CYP3A4 may overlap (Harlow and Halpert, 1997). Also, Leu210
and Leu211 of CYP3A4 are highly conserved between species; however,
CYP3A5 has Phe at residue 210. The Leu210 Phe substitution did not
alter the enantiomer ratio (1.06). This finding suggests that the
difference in the ratio of sulfoxide enantiomer formation between
CYP3A4 and 3A5 cannot be accounted for by residue 210, a conclusion
that is consistent with the 3A4 modeling data.
The residues that encompass SRS-4 are highly conserved between species
and have been directly implicated in substrate specificity and
metabolism (Szklarz and Halpert, 1997; Fukuda et al., 1993; Raag and Poulos, 1989). Of the four SRS-4 mutants tested, only T309A
showed sulfoxidase activity comparable with that of the wild type.
F304A and I301A showed decreases only in R-sulfoxide formation. Interestingly, F304A has previously shown greater
progesterone hydroxylase activity in comparison to the wild type,
suggesting that the mutation does not effect the inherent function of
the enzyme, but more likely the binding of the particular substrate. This interpretation is supported by the prediction of a distance of
only 4 Å between a docked progesterone molecule and the 304 residue
(Szklarz and Halpert, 1997). Residue T309 also has been mapped to close
proximity to CYP3A4 substrates and has also been implicated in the CYP
catalytic cycle, a finding supported by the change in enzyme activity
with the mutation of the aligned residue of rabbit 2E1 and rat 2A1
(T303; Fukuda et al., 1993). However, steroid 6-hydroxylation (B
ring of the steroid) and now RPR 106541 sulfoxidation (D ring
substituent) have been shown to be unaltered by the threonine to
alanine change at residue 309, thus highlighting the importance of
model validation with metabolism data.
Steroid 6-hydroxylation by CYP3A4 has also been shown to be altered
by the mutation of residues I369, A370, and L373 (He et al., 1997).
With RPR 106541 sulfoxidase activity, however, only the A370V mutation
resulted in a dramatic change in the rate of metabolism and the
enantiomeric ratio (0.23). The A370V substitution has been implicated
in the stabilization of progesterone in the 16 binding orientation
(D ring as opposed to the 6 or B ring orientation), and may explain
why the A370V mutant had the highest rate of S-sulfoxide
formation of any mutant (Szklarz and Halpert, 1997). The effect of
alterations of SRS-6 residues on steroid hydroxylase activity for CYP2B
forms has been shown to be dramatic. Substrate docking studies have
shown that residues 478-480 map to SRS-6 and potentially interact with
the substrates erythromycin and progesterone (Szklarz and Halpert,
1997). Consistent with the docked substrate model, substitution of a
Phe for L479 selectively decreased R-sulfoxide formation.
The fact that the ratio of 6-OH and 16-OH metabolites of
progesterone and testosterone were not effected by this mutation
illustrates the increased sensitivity of RPR 106541 to SRS-6 mutations.
In conclusion, we have shown that the use of homology modeling in
combination with site-directed mutagenesis can add important information on CYP-450-substrate interactions to CYP reaction phenotyping studies. There are three reasons why CYP3A4 is an excellent
candidate for studies that could potentially advance the descriptive
phenotyping data of individual compounds to more predictive models of
CYP3A4-substrate interactions. First, CYP3A forms are responsible for
the majority of xenobiotic oxidations in humans, with CYP3A4
representing the largest percentage of CYP forms in liver and a
significant barrier to bioavailability in the intestine. Second, CYP3A4
displays a relatively high homology (88%) to CYP3A5, a polymorphically
expressed form whose levels can equal or exceed those of CYP3A4 in
certain individuals. This amino acid sequence similarity could be
exploited to narrow the search for differences in CYP3A4/3A5 substrate
interactions and provide critical information on interindividual
variability in the metabolism of compounds with overlapping CYP3A
substrate specificity. Finally, there is a large literature base on the
stimulation of CYP3A4-catalyzed reactions in liver microsomes and, more
recently, in human hepatocytes (Maenpaa et al., 1998). The combination
of homology modeling with site-directed mutagenesis may also help to
differentiate a number of hypotheses proposed on the mechanism of CYP3A
stimulation (Schwab et al., 1988; Shou et al., 1994; Ueng et al., 1997;
Harlow and Halpert, 1998). The historical anomalies associated with
many CYP3A4-catalyzed reactions represent a growing research area that
can now be addressed by a variety of substrates and genetic engineering techniques.
We acknowledge You-Ai He for kindly providing several purified
mutants, Dr. Grazyna Szklarz for providing the CYP3A4 model, Dr.
Sheng-Yuh Tang for LC/MS support, and Dr. Andrew Bridge for the
synthesis of metabolite standards.
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Abstract
Introduction
